Celluar Machanisms of Insect Photoreception

INTERNATIONAL REVIEW OF CYTOLOGY,VOL. 57

Cellular Mechanisms of Insect Photoreception F. G. GRIBAKIN The Laboratory of Evolutionary Morphology, Sechenov Institute of Evolutionary Physiology and Biochemistry, Academy of Sciences of the USSR, Leningrad. USSR

I. Introduction . . . . . . . . . . . . . . . . . . . . 11. The Compound Eye and Photoreceptor Optics . . . . . . . . A. Light and the Eye . . . . . . . . . . . . . . . . B. The Compound Eye . . . . . . . . . . . . . . . . C. Visual Pigments and the Photoreceptor Membrane . . . . . D. Absolute Light Sensitivity of the Compound Eye . . . . . E. Color Vision . . . . . . . . . . . . . . . . . . F. Sensitivity to Polarized Light . . . . . . . . . . . . G. Final Remarks . . . . . . . . . . . . . . . . . . 111. Electrical Basis for Insect Photoreception . . . . . . . . . A. Introduction: The Compound Eye as a Volume Conductor . . B. Dc Parameters of the Compound Eye in the Dark and in the Light C. Photoresponses and Their Cellular Mechanisms . . . . . . D. The Role of Compartmentalization . . . . . . . . . . IV. Conclusion . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . . .

127 128 128 129 129 138 153 157 158 159 159 159 169 177 177 178 184

When you go after honey with a balloon, the great thing is not to let the bees know you’re coming. Now, if you have a green balloon, they might think you were only part of the tree, and not notice you, and if you have a blue balloon, they might think you were only part of the sky, and not notice you, and the question is: Which is most likely? A. A. Milne, Winnie-the-Pooh

I. Introduction The photoreceptor of insects is a highly specialized sensory cell whose principal function is to transform information on the arrival of a light quantum into a receptor signal. At the same time, all the specific machinery of the insect photoreceptor is undoubtedly based on general cellular mechanisms underlying not 127

Copyright Q 1979 by Academic Press, Inc. All rights of reproduclion in any form reserved. ISBN 0-12-364357-0

128

F. G. GRIBAKIN

only sensory cell functioning but, to some extent, the functioning of a neuron. However, unlike the neuron, which has many neuronal (or synaptic) inputs, the photoreceptor cell has only one input which is physical. Such a peculiarity of the photoreceptor has determined the logic of this article which discusses the cellular mechanisms of photoreception in insects from the absorption of a light quantum to generation of the receptor signal. Both the start and the end of the insect photoreceptor process have been studied fairly well, however, many intermediate stages are still unknown, and, we will emphasize those aspects of the problem we believe to be of paramount importance, at least, at the end of the 1970s. We apologize to those whose original works are not referred to in this article, since the number of references had to be reduced to a reasonable minimum. This article is addressed not only to specialists in cell biology but also to investigators dealing with the nervous system, since the compound eye of arthropods can serve as an excellent model for various neurophysiological studies. 11. The Compound Eye and Photoreceptor Optics

A. LIGHTAND THE EYE Undoubtedly, all the morphological and physiological characteristics of the eye, as well as of its photoreceptors, have been totally predetermined by the nature of light. As an electromagnetic wave, light can be characterized by energy (or intensity), wavelength, polarization, and direction of propagation. A photoreceptor must display absolute, spectral, polarization, and directional sensitivities which at the whole-eye level are known as light sensitivity, color vision, polarized light discrimination, and image perception. Investigation of these characteristics, based on the fundamentals of cell biology (cytology) and physical optics, forms the subject of a relatively new branch of science called photoreceptor optics, the birth of which was recently announced at a special workshop by Menzel and Snyder (1975). Here we consider the absolute, spectral, and polarization sensitivities of the photoreceptors of the photopic compound eye in terms of photoreceptor optics. Directional sensitivity and image perception have been omitted from this account, since these subjects are closely related not to cell biology but rather to three-dimensional space perception, a field in which we have had no personal experience. The term “scotopic” and “photopic” are used here instead of “superposition” and “apposition,” since we accepted terminology of Post and Goldsmith (1965) who suggested to discriminate between compound eyes with short proximal rhabdoms and migratory pigment (scotopic) and those with long rhabdoms and no longitudinally migrating pigments (photopic).

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

129

B. THECOMFQUND EYE The compound eye consists of structural units called ommatidia. The three main elements of an ommatidium are the dioptric apparatus, retinula, and pigment sheath. In turn, the dioptric apparatus includes a corneal lens (the biconvex transparent cuticule region) and a crystalline cone produced by four transparent cone cells. Nearly round in cross section, the retinula is formed by several (usually eight) visual cells or photoreceptors, each producing a membraneous photoreceptive structure termed a rhabdomere. In many insects rhabdomeres fuse axially in the retinula to form a rhabdom. A photoreceptor sends its central process, or axon, through the basal membrane (or basal lamina) toward higher nerve centers-optic ganglia (the lamina, medulla, and lobula). The pigmented sheath of an ommatidium is composed of primary (or comeagenous or iris), secondary (or accessory), and sometimes basal pigment cells. It should be emphasized that the compound eye is encapsulated within chitinous walls and proximally separated from underlying structures and optic centers by a basement lamina. Thus the extracellular space of the compound eye should be considered an extracellular compartment isolated from the rest of the insect (Heisenberg, 1971). We omit further description of the compound eye structure and classification, as well as ultrastructural details, since they can be easily found in the literature (see, for review, Mazokhin-Porshnyakov, 1969; Goldsmith and Bernard, 1974). C. VISUAL PIGMENTS AND THE PHOTORECEPTOR MEMBRANE 1. Introduction Since the retina is intended to absorb light, it cannot be as transparent as common nerve tissue. The stronger its coloration, the greater the amount of light absorbed and the greater its absolute sensitivity. Dealing with photoreception, natural selection might realize four ways to get the photoreceptor cell colored; it might stain (1) cell cytoplasm, (2) intracellular inclusions and organelles, (3) the plasma membrane, or (4) all three. Thanks to the efforts of many workers, most of all G. Wald, we know that the third possibility has been realized in vision, and that the absorbing substance of a photoreceptor-the visual pigment-is an integral part of specialized areas of the outer, or plasma, membrane of the cell, which therefore has been called the photoreceptor membrane. Current knowledge of the photoreceptor membrane derives mainly from studies on the retina of vertebrates. However, all we know about the arthropod photoreceptor membrane indicates that both have much in common. Thus, for simplicity, we discuss the visual pigments and optical properties of these membranes as though they were the same "universal" photoreceptor membrane (Gribakin and Govardovskii, 1975), and particular emphasis is placed upon their differences when appropriate.

130

F. G . GRIBAKIN

2. Visual Pigments a. Retinal as a Chromophoric Group in Insect Rhodopsin. A visual pigment molecule is believed to consist of a lipoprotein moiety to which a chromophoric group is attached. The first visual pigment studied in detail was rhodopsin extracted from retinal rods of cattle and frogs and having a maximal absorption A,,, at 500 nm. At present all visual pigments are usually termed rhodopsins, no matter what class of animals they belong to or where their A,, is located. The basis for this generalizationis the chromophoric group 11 &-retinal (a derivative of pcarotene) which is the same in almost all pigments known; only porphyropsins found in some vertebrates, which have 3,3’-dehydroretinal as a chromophoric group, are exceptions to this rule. In dark-adapted vertebrate photoreceptors the chromophoric group is bound to the phospholipid phosphatidylethanolamine (Abrahamson and Wiesenfeld, 1972). After absorption of a light quantum and subsequent cis-trans isomerization, the chromophoric group is transferred from the lipid to a lysine group on a protein (opsin), and this coincides with the change from metarhodopsin I (A,, = 478 nm) to metarhodopsin I1 (A,, = 380 nm). Of all the intermediate transitions (which occur in the dark and at room temperature), this is the first to require water, and it is thought that the metarhodopsin transition gives rise to the generation of a late receptor potential, or the receptor signal (Abrahamson and Wiesenfeld, 1972). This is not true for cephalopod rhodopsins where the retinal binding site is located on a protein, and light-evoked transitions stop at a stage of metarhodopsin formation (Hubbard and St. George, 1958). It means that, unlike the final product of vertebrate rhodopsin transitions-N-retinylidene opsin, which readily hydrolyzes to yield free retinal and opsin (so-called bleaching of the pigment)-cephalopod metarhodopsin is resistant to hydrolysis (for more details, see Abrahamson and Wiesenfeld, 1972). Further, since in cephalopods water is involved in the lumirhodopsin-metarhodopsin transition, this stage may be considered an analog of the vertebrate metarhodopsin I-metarhodopsin I1 reaction responsible for initiation of the receptor signal. Visual pigments of arthropods resemble cephalopod rhodopsins rather than those of the vertebrate retina in having stable or long-lived metarhodopsins (see, for review, Goldsmith, 1972). According to the evolutionary classification of Eakin (1965, 1972), both cephalopod molluscs and arthropods have rhabdomeric photoreceptors, so the similarity in photochemistry may reflect a fundamental property of the rhabdomeric photoreceptor. The small dimensions of the insect compound eye make it difficult to investigate its visual pigments in vitro. Nevertheless, Goldsmith (1958) was the first to show that retinal occurred in honeybee heads but not in abdomens and thoraxes. The same proved to be true for six species of other orders: Hymenoptera, Orthoptera, Odonata, Coleoptera, and Lepidoptera (Briggs, 1961). Retinal has also been found in Diptera (Wolken et al., 1960) and Blattoptera (Wolken and Scheer, 1963). Free retinol has been

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

131

reported in light-adapted bees, although retinal usually appears to prevail in a state of dark adaptation (Goldsmith and Warner, 1964). The absolute sensitivity of the eye of carotene-depleted flies (Musca dornestica) has been found to drop by about four orders of magnitude in both the visible and ultraviolet regions possibly because of a shortage of chromophoric substance-retinal (Goldsmith et al., 1964; Goldsmith and Fernandez, 1966). Paulsen and Schwemer (1972) demonstrateddirectly that retinal in the 1 1 -cis form was actually the chromophoric group of Ascalaphus macaronius ultraviolet-sensitive rhodopsin-the first visual pigment of insects isolated and photochemically characterized (for its history, see Gogala and Michieli, 1965; Gogala, 1967; Gogala et al., 1970; Hamdorf et al., 1971; Schwemer et al., 1971). Thus there is ample reason to extend the above findings to the whole class of insects and to suggest that all insect rhodopsins incorporate 11 -cis-retinal (never 3-3’dehydroretinal) as a chromophoric group. b. Absorption Spectra of Znsect Rhodopsins. In vertebrates visual pigments incorporating 11-cis-retinal range in A,, from 433 to 575 nm (Dartnall and Lythgoe, 1965; Liebman and Entine, 1968). Between these points the absorption maxima of about 200 known vertebrate rhodopsins are not randomly distributed, but cluster about certain points of the spectrum about 7 nm apart (Dartnall and Lythgoe, 1965). As judged from in vivo spectrophotometry and, mainly, electrophysiology, insect rhodopsins show a greater diversity of A,, values (Goldsmith and Bernard, 1974; Menzel, 1975b). The red limit for A,, in insects does not seem to exceed the 575 nm reported for vertebrates (e.g., 567 nm in the green receptors of Notonecta, Bruckmoser, 1968), and the presence of redsensitive pigments can not be inferred as yet from the data available from electrophysiology (Goldsmith, 1965; Swihart, 1972; Post and Goldsmith, 1969; Autrum and Kolb, 1968; also see review of Menzel, 1975b). The most distinctive feature of insect rhodopsins (as well as of arthropod rhodopsins in general) is the presence of retinal-based pigments with a A,,, in the near ultraviolet region (340-360 nm) and with practically no absorption in the visible region. Theory predicts both bathochromic (to the longwave side) and hypsochromic (to the shortwave side) shifts of the A,,, from 435 to 440 nm inherent in essential visual pigment of vertebrates ( N - 1 1-cis-retinylidene phosphatidylethanolamine). These may result from twisting about a double bond in the chromophore, which depends upon the molecular microenvironment of the chromophore, or from interaction of the chromophore with a closely located negative charge (Abrahamson and Wiesenfeld, 1972). It appears, however, that even twisting cannot explain the full range of A,, values reported in vertebrates (433-575 nm) let alone in the ultraviolet region, whereas a model calculation of the effect produced by a negative point charge placed 0.45 nm above the plane of that are both bathothe chromophore molecule demonstrates shifts in A,, chromic (to 530 nm when positioned at the fifth carbon of retinal) and hypsochromic (to 356 nm when displaced toward the nitrogen atom of the binding site)

132

F. G. GRIBAKIN

(see Table 6 in Abrahamson and Wiesenfeld, 1972). An alternative possibility is the linkage of retinal with the sulfhydryl group of cysteine, or with both the sulfhydryl and amino groups of opsin (Goldsmith, 1972). Thus the existence of retinal-based ultraviolet-sensitive rhodopsins may be postulated from theory (see also Morton and Pitt, 1969). All the data available on color receptors in insects are presented in Fig. 1. Although their A,, values may not coincide exactly with those for their rhodopsins for many reasons (Section 11,E; Menzel, 1975b), we can infer that longwave (green and blue) receptors of insects located within the same broad spectral band as vertebrate retinal-based photoreceptors do (412-567 nm in insects versus 433-575 nm in vertebrates). Only 2 of about 50 color receptors have sensitivity peaks between 380 and 420 nm, which may imply an “exclusion principle” for this region, and, in turn, this may have a real physical or physiological basis. As judged from Fig. 1, visual pigments of insects tend to cluster around four distinct points of the spectrum, 350, 430, 470, and 520 nm; the reason for this is still unknown. c. Energetics of the Photoexcitation and Regeneration of Insect Rhodopsin. Visual pigments of animals originate from p-carotene synthesized in plants. But, unlike the situation in plants, in which the energy of light absorbed by chlorophyll, carotenoids, or accessory pigments is used metabolically, in animals the energy of a light quantum absorbed by a rhodopsin molecule does not appear to serve as the energy supply for further stages of the visual process. The only function of a light quantum in vision is to release from the photoreceptor cell an appropriate “quantum” of metabolic energy which has been stored in it as electricity. Thus the molecular evolution of visual pigments has changed the light quantum function from that of an energy supplier in plants to that of an information carrier in animals (Vinnikov, 1974).

iII N=50

400

5

I

500

,

600 A , nm

FIG. 1 . Distribution of insect color receptors over the spectrum. (Maxima positions are mainly from Goldsmith and Bernard, 1974.)

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

133

A molecule of bovine rhodopsin-498, upon absorption of a light quantum, transforms through an intermediate-prelumirhodopsin-543 (still with an 11-cis chromophoretinto lumirhodopsin-497 (with an all-trans chromophore) (Morton, 1972). The energy levels of both rhodopsin-498 and lumirhodopsin497 are nearly the same (see, for instance, Abrahamson and Wiesenfeld, 1972), so the energy of a light quantum is necessary only for the molecule to overcome the potential barrier between the two states encoding either the absence of light (rhodopsin) or the presence of light (lumirhodopsin). All the excessive energy liberated during the “fall” of the molecule from the potential barrier to the lumirhodopsin state dissipates as heat (see discussion in Falk and Fatt, 1972). The other mechanisms which could help the molecule to dispose of the unnecessary energy do not seem to be of any significance (e.g., the quantum yield of the fluorescenceof rhodopsin is as small as 0.005; see Guzzo and Pool, 1968). Thus, after absorption of a light quantum (photoexcitation) followed by cis-trans isomerization (photoisomerization), the rhodopsin molecule, which has already triggered a mechanism for signal generation, leaves the field through several spontaneous (thermal) transitions ending with its breakdown into free retinal and opsin. In order to keep the absolute sensitivity of the photoreceptor high enough, a continuous renewal, or resynthesis, of rhodopsin molecules is needed, and two methods of regenerating rhodopsin are known-enzymic and photochemical. Enzymic regeneration is most characteristic of vertebrates. However, of all the vertebrates studied, only the rat seems to have an effective isomerizing system in the retina (Cone and Brown, 1969). In other vertebrates (frogs, cattle) isomerizing systems are thought to be located mainly in pigment epithelium, where 11-cis-retinol esters dominate other retinol isomeric derivatives (see, for review, Baumann, 1972). Photoregeneration of rhodopsin in vertebrates can be demonstrated in three ways. First, strong illumination or a brilliant flash is used so that the photoexcited molecule of rhodopsin can absorb a second quantum during its short stay in the lumi- or metarhodopsin state, that is, before its thermal breakdown begins. The second quantum thus absorbed can convert the intermediate molecule into its native state-rhodopsin-or into isorhodopsin with 9cis-retinal as a chromophore [see the experiments of Dowling and Hubbard (1963) with the rat retina]. Second, the temperature can be lowered so as to stablize the intermediate. In this case prolonged illumination of rhodopsin preparations produces an equilibrium mixture composed of 11-cis, 9-cis, and all-trans modifications of visual pigment (Yoshizawa, 1972). Third, shortwave light, blue or violet, with its wavelength lying within the absorption band of all-trans-retinal can be used. Under these conditions, light bleaches rhodopsin, but all-trunsretinal liberated from bleached pigment undergoes photoisomerization resulting in 11-cis isomer formation, and the latter can combine freely with opsin. Thus, partial regeneration of rhodopsin (to 15-20%) is again possible (Hubbard and Wald, 1952). From this consideration one can infer that photoregeneration of

134

F. G. GRIBAKIN

rhodopsin in vertebrates demands very specific light or temperature conditions and, for this reason, can hardly be of any importance in their visual cycle. On the contrary, in insects as well as in other arthropods, and also in cephalopod molluscs (all having rhabdomeric eyes), photoregeneration prevails in the restoration of visual pigments. As mentioned above, light-evoked changes in rhodopsins in these groups result in metarhodopsins (Hubbard and St. George, 1958, for the squid; Hamdorf et al., 1968, for the octopus; Wald, 1967, for the crayfish; Gogala et al., 1970; Hamdorf et al., 1971; Schwemer et al., 1971, for the insect Ascalaphus). The same appears to be true for Calliphora, Drosophila, and Deilephila, and in all these species photoregeneration of rhodopsin from long-lived or thermally stable metarhodopsin has been demonstrated (see, for review, Hamdorf and Schwemer, 1975). Since the absorption coefficients of both rhodopsin and metarhodopsin yR(A) and yM(A), respectively, depend on the wavelength of the illumination, absolute quantities of both pigments in the photoreceptor depend only on wavelength and quantum yields of direct and reverse photoreactions. Assuming quantum yields to be equal and illumination prolonged enough, one can obtain a rhodopsidmetarhodopsin ratio at a given Ao, as [R]/[M] yM(AO)/yR(Ao), which results in a maximal rhodopsin content when A. corresponds to the A,, of the metarhodopsin absorption spectrum, and in a maximal metarhodopsin content when A. is the A,, of the rhodopsin absorption spectrum. Thus prolonged illumination with monochromatic light (under experimental conditions) or wide-band light (in a natural environment) inevitably leads to a photoequilibrium between the rhodopsin and metarhodopsin in the photoreceptor. As experiments with Deilephila elpenor (Lepidoptera) have shown, the same metarhodopsin (M480) forms from different rhodopsins (R350, R450, and R545) in different color receptors of the same eye (Schwemer and Paulsen, 1973). There is evidence that the energy of a light quantum absorbed by metarhodopsin is needed only to convert all-trans-retinal to 1 1-cis-retinal but is not necessary to produce reverse conformational changes in the protein from the metarhodopsin to the rhodopsin state (Hamdorf and Schwemer, 1975). Also, metarhodopsin, no matter what its amount in the photoreceptor, has no influence on signal transduction which is totally defined by rhodopsin function and amount (Hamdorf and Schwemer, 1975), although it may affect the optical properties of the photoreceptor by optical screening of rhodopsin. Dark regeneration of rhodopsin (probably enzymic) from metarhodopsin has also been shown to occur in photoreceptors of invertebrates, although this process is slow. So, to convert half the initial amount of metarhodopsin into rhodopsin about 10 minutes at 20°C is needed in the octopus Eledone moschata (M520 to R470, Schwemer, 1969), about 25 minutes at room temperature in Calliphora (M580 to R495, Stavenga et al., 1973), 15-45 minutes at room temperature in the butterfly Aglais urticae (M480 to R535, Stavenga, 1975), and several minutes in the lobster Homarus (M490 to R515, Goldsmith and Bruno, 1973). At the

-

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

135

same time, Brown and White (1972) failed to find dark regeneration of rhodopsin R515 from M480 in larvae of the mosquito Aedes aegypti. Thus photoregeneration of visual pigments seems to play a leading part in the insect visual cycle (as well as in the visual cycles of other invertebrates), whereas dark (or metabolic or enzymic) regeneration is probably a continuously working standby mechanism which, although it is not so powerful as photoregeneration, could activate the visual system into a “ready-to-see’’ state under poor light conditions or even in the dark when photoregeneration is impossible (cf. Stavenga, 1975). 3 . The Photoreceptor Membrane In 1962 G. Wald formulated the concept of a photoreceptor membrane: “It is at least very reasonable to expect that porphyropsin . . . is located in the membranes” (Wald et al., 1962). Since photoreceptor membranes of different species have much in common, the variety of light-sensitive organelles known in the animal kingdom can be considered as having evolved from the originally flat universal photoreceptor membrane (Gribakin and Govardovskii, 1975; Laughlin et al., 1975; Menzel and Snyder, 1975). Rhodopsin molecules are inlaid in this membrane, and in vertebrates they are free to undergo Brownian rotation and translation (Brown, 1972; Cone, 1972; Po0 and Cone, 1974; Liebman and Entine, 1974). Thus the photoreceptor membrane conforms to the model of a fluid mosaic membrane with protein molecules floating in a lipid bilayer (cf. Singer and Nicholson, 1972; Edidin, 1974). The rod outer segment may be considered a “pure preparation” of photoreceptor membrane, and its lipid and protein content well reflects the composition of the photoreceptor membrane. The lipid content is 40% of the total dry weight of the rod outer segment, and proteins comprise about 60%. The protein moiety of rhodopsin-opsin-constitutes up to 80-90% of the total protein content in the frog rod outer segment and 15-20% in cattle (Abrahamson and Wiesenfeld, 1972). Rhodopsin molecules are believed to be located in the inner (facing the cytoplasm) lipid layer of the photoreceptor membrane, that is, on the outside in rod disks and on the inside in rhabdomeric microvilli (Blasie, 1972; Daeman, 1973; Mason et al., 1974; Jan and Revel, 1974, for outer segments; Hamdorf and Schwemer, 1975; Fernandez and Nickel, 1976; Eguchi and Waterman, 1976; Nickel and Menzel, 1976, for rhabdomeres). The surface concentration of rhodopsin in outer segments is about (2 X 1 0 4 ) / ~ m z (a “square lattice” with molecules 7 nm apart; see, for instance, Daeman, 1973); calculated for a crayfish rhabdomere this is about half less, that is, (1 x 104)/pm2 [a 9 x 9 nm square lattice as calculated by Gribakin and Govardovskii (1975) from the data of Hays and Goldsmith (1969)l. Freeze-etch study of the crayfish rhabdomeric membrane gives a similar surface density of protein particles on its inner side-about 8000/pm2 (Eguchi and Waterman, 1976). Nearly the same value, 7000/,um2, has been obtained for the ant (Nickel and Menzel, 1976) and

136

F. G. GlUBAKIN

the crayfish Procamburus (6600/pm2, Fernandez and Nickel, 1976). There is much evidence that these particles correspond to rhodopsin molecules in vertebrate photoreceptors (see Jan and Revel, 1974). This is also thought to be true for the rhabdomeric membrane (Fernandez and Nickel, 1976; Nickel and Menzel, 1976), and the change in surface density of particles observed in carotenedepleted flies strongly supports this view because of the numerical coincidence between the decrease in rhodopsin content (about eight times) and in the surface density of particles (nearly six times) (Boschek and Hamdorf, 1976). The diameter of a vertebrate rhodopsin molecule (with a molecular weight of 40,000 daltons) has been suggested to be 4-5 nm (Wald er al., 1962), and the globular substructures in the disk membrane appear to have the same size (Daeman, 1973). This implies that at above surface concentrations [(2 x lo4)/ pm2] about 60% of the membrane surface is free of rhodopsin (it would be only 21 or 14% for square or hexagonal packing). So at this concentration there is no need for rhodopsin molecules to be packed in any kind of crystalline lattice; instead, they are seemingly free to locate at random. The larger diameter of particles in the invertebrate eye membrane-mean 8 nm (Eakin and Brandenburger, 1975, for the snail; Fernandez and Nickel, 1976, for the crayfish; Nickel and Menzel, 1976, for the anttsuggests a molecular weight of the pigment as high as 200,000 daltons, and only if the rhodopsin molecule in the particle were surrounded by two layers of lipids could this value be lowered to 52,000 daltons, close to that of vertebrate rhodopsins (Nickel and Menzel, 1976). Another explanation of the large size of particles suggests that the rhodopsin molecule is a dimer, or even a multimer (Nickel and Menzel, 1976), however, this is not consistent with optics from which a surface density of absorbing centers (i.e., rhodopsin chromophores) of about (1 X 104)/pm2has been obtained (see above), while a hypothetical dimer or multimer would have shown absorption two or several times greater than that measured by Hays and Goldsmith (1969). Vertebrate rhodopsin molecules are free to rotate and translate, but strict limitations exist as to spatial orientation of their rotation axes so that tumbling “head over heels” is highly improbable. Owing to these restrictions, the rhodopsin molecule behaves as if it were suspended in its molecular environment from two points-a hydrophilic “top” jutting out into the aqueous phase (cytoplasm) and a hydrophobic “bottom” embedded in the lipid phase of the membrane (cf. Singer and Nicholson, 1972; Laughlin et al., 1975). This rotational suspension of the molecule in the photoreceptor membrane can be considered one of the principal attainments in the molecular evolution of rhodopsin which made it possible to orient all the molecules (about 3 x 10’ in the frog rod outer segment, for instance) in a standard position providing maximal absorption of the light normally incident upon a flat membrane. [Maximal and isotropic absorption results from keeping the absorbing dipoles of properly suspended rhodopsin molecules at a right angle to the suspension axes, i.e., in a plane of the photoreceptor membrane, a fact first discovered by Schmidt (1938), Wald er al.

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

137

(1962), and Liebman (1962).] Rolling up of the flat universal membrane, which is necessary to form a microvillus, puts additional restrictions on rhodopsin Brownian translation (Wehner and Goldsmith, 1975) and rotation (Goldsmith and Wehner, 1975; Laughlin et al., 1975). Using microspectrophotomet, Wehner and Goldsmith (1975) observed no translational diffusion of metarhodopsin produced by a light flash in crayfish photoreceptor membranes. In addition, the absorbing dipoles of rhodopsin are partially aligned along the axes of microvilli in the squid and crayfish (microspectrophotometry, Hagins and Liebman, 1963; Goldsmith, 1975), which follows from the dichroic ratio of a microvillus, which is about 2.5-2.7 (mean value) in the crayfish and 6 in the squid, instead of the expected 2.0 if the ideal dipoles were oriented at the plane of the membrane at random (the model of Moody and Parriss, 1961). Several physical reasons responsible for the above restrictions were put forward. First, a lower proportion of long-chain polyunsaturated fatty acids with more than 20 carbons (27.3% of the total fatty acids in squid rhabdoms versus 42% in vertebrate rods), as well as a higher amount of cholesterol (with a molar ratio to phospholipids in the squid of about 1:2 versus 1:20 in vertebrate rods), make the rhabdomeric membrane more rigid or stiff, and consequently more viscous (Goldsmith, 1975; Hamdorf and Schwemer, 1975). Second, since the surface of a microvillus is curved, a boundary between different phases of the membrane (hydrophilic and hydrophobic) would be disturbed periodically during rhodopsin molecule rotation, and this would be energetically unfavorable. Thus bending of the membrane might prevent molecules from free rotation in the membrane, though oscillations or wobbling may occur (Goldsmith and Wehner, 1975; Laughlin et al., 1975). Third, the molecule may be asymmetric in the plane of the membrane, hence if subjected to viscous drag (e.g., from hypothetical cytoplasmic flows within the microvillus; see Goldsmith, 1975), it might move or oscillate with its long axis directed along the microvillus (Laughlin et al., 1975). Fourth, curvature of the microvillar membrane seems to prevent circumferential translation energetically (Laughlin et al., 1975). Fifth, taking into account the location of rhodopsin molecules inside the microvillus, the calculated area per molecule can be lowered from about 80 nm2 (for an outer diameter of the microvillus of 50 nm) to 65 nm' (for an inner diameter of about 40 nm). Then, for 8-nm particles one can obtain a surface density closely corresponding to a square lattice with little room for free diffusion. Last, an interaction between closely positioned molecules may occur which could stabilize the positions of the molecules and eventually activate the microvillar photoreceptor membrane into a state similar to the crystalline state. Most of the above restrictions may apply to a greater extent in microvilli of smaller diameter (Laughlin et al., 1975). Thus the rhabdomeric photoreceptor membrane has evolved to incorporate rhodopsin molecules in more-or-less fixed positions, and this arrangement seems to be a consequence of the bending (or rolling up) of an originally flat photo-

138

F. G. GRIBAKIN

receptor membrane to form a tubule or microvillus. In this situation photoregeneration has proved to be the more universal way of restoring rhodopsin, because of the easy accessibility of any molecule to light, while it does not seem that natural selection could easily supply every rhodopsin molecule fixed in the membrane with a quickly operating enzymic regeneration mechanism of its own.

D. ABSOLUTE LIGHTSENSITIVITY OF THE COMPOUND EYE 1. Principal Problems In order to design an ideal photoreceptor evolution (or natural selection) had to solve at least two principal problems: first, how to transduce information on every photon absorbed to the proper discrete receptor signal (one-to-one transduction) and, second, how to maximize the ability of a receptor to absorb photons. The first problem is known to have been successfully solved both in vertebrate and invertebrate photoreceptors which can act as photon counters (Hecht et al., 1942; Sakitt, 1972, for humans; Fuortes and Yeandle, 1964; Scholes, 1965; Borsellino and Fuortes, 1968; Kaplan and Barlow, 1976, for arthropods). The efficiency of counting is limited by the quantum yield of rhodopsin which, being 0.67 in vertebrates (see Dartnall, 1972), means that one light quantum from the three absorbed fails to evoke bleaching and apparently the unit electrical response. In arthropods, unit electrical responses were first described by Yeandle (1958) in Lirnulus, and at present they are known as quantum “bumps,” discrete potentials, or miniature potentials in all the arthropod species studied. So, it appears that the first general problem of vision-one-to-one transduction from a quantum of light to a “quantum of electricity”-was solved early in evolution and that since then the second problem has appeared-how to increase the probability of absorption of a photon. Unlike a one-to-one transduction mechanism which must be considered in terms of membrane physiology (Section III,C), the evolution of light absorption by photoreceptors is totally within the sphere of photoreceptor optics.

2. The Beer-Lambert Law as Applied to Rhabdomeric Photoreceptors Quantitatively transmission of light (and consequently its absorption) is expressed by the well-known Beer-Lambert law for an absorbing layer: Zt = Zo exp

-

a (A, 40) cl

(1)

where It and Zo are the intensities of the transmitted and incident light, respectively, a (A, cp) is the extinction (absorbance)coefficient depending in general on the wavelength A and the angle cp between the E vector of t b incident wave and the axes of the absorbing dipoles, c is the concentration of the absorbing substance (pigment), and I is the path length, or the thickness of the absorbing layer. Equation (1) can be written in another form using Napierian logarithms:

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

139

or, using common logarithms, = log,o(~t/~o)

E

(A, cp)cl

(3)

where, for 1 in centimeters and c in moles per liter, E (A, cp) has dimensions of liters per centimeter-mole and is termed the molar extinction (or molar absorbance). The optical density (or absorbance) D is often used, since it is more convenient:

D

= -log&/&)

=

E

(A, cp)d

(4)

The greater D, the greater the absorption, so in order to increase the absolute sensitivity of a photoreceptor (i.e., to maximize the probability of photon absorption), all three factors, namely, E (A, cp), c, and 1 must be made as large as possible. a. Molecular Extinction Coeficient, Molar Extinction, and the Orientation of Rhodopsin Molecules. If the path length 1 is expressed in centimeters and the concentration c in number of molecules per cubic centimeter, a (A, cp) is the molecular extinction coefficient having the dimensions of area. a(A, cp) and the molar extinction E (A, cp) are connected by a simple relation:

(A, cp) = (2.303

X

103) [ E (A,

cp)/Nl

(5)

where N is Avogadro’s number. In terms of physics, a (A, cp) is related to the effective area of the absorbing site of the molecule multiplied by the probability that a photon striking the molecule within this area will be absorbed. The change in this probability with wavelength forms the natural basis for the absorption spectrum, while its dependence upon the angle cp reflects dichroic properties of the molecule. At the maximum of its absorption spectrum (Amax) randomly oriented vertebrate rhodopsin (e.g., in a solution) has an average (Y (A,,, p) of 1.56 x cm2 (Dartnall, 1972). For linearly polarized light with A = A,, and (c = 0 or cp = T (when all the absorbing dipoles are parallel to the E vector), the highest value of the molecular extinction coefficient is three times greater, that is, 4.68 x cm2, since only one-third of the dipoles were parallel to the E vector when measured in a solution. This situation resembles that reported for rhabdomeric membranes with predominant dipole alignment along the microvilli. However, for unpolarized light, half the uniformly oriented dipoles make no contribution to absorption and, in order to obtain maximal absorption in this case, the dipoles should be allowed to arrange at random (or to rotate) in the plane perpendicular to the light path; this has been found to be true for vertebrate rod disks. The above considerations are illustrated by Fig. 2 and Table I where four theoretically possible modes of orientation are given. Several important conclusions can be made based on these models. First, the value of a (A, cp) for

140

F. G . GRIBAKIN

Y/

FIG.2. Scheme illustrating the dependence of the average molar extinction of rhodopsin on the orientation of absorbing dipoles. A unit volume is filled with a rhodopsin solution with concentration c and partial concentrations c, c , , and c , ; I, is the intensity of the incident light; E, and E, are the components of the E vector. Self-screening has been neglected. Quantitative analysis is given in Table I.

rhodopsin measured in solution is three times less than the maximal value. The latter is the most appropriate parameter of the absorbing dipole when one deals with pigment orientation in the photoreceptor membrane (4.68 X cm2 or emax= 12.18 x lo4 literkm-mole; cf. 1.56 X cm2 and emax = 4.06 x lo4 literkm-mole when in solution; see Dartnall, 1972). Second, twodimensional orientation of absorbing dipoles (“disks”) produces a 1.5-fold increase in the average molar extinction (as compared to that measured in solution) for both polarized and unpolarized light. Such a system fails to discriminate between polarized and unpolarized light when light travels normal to the disk plane. Third, an all-parallel mode of orientation of absorbing dipoles gives maximally a 3-fold gain in the average extinction coefficient for polarized light and a 1S-fold gain for unpolarized light. Thus all-parallel orientation gives no gain in absolute sensitivity to unpolarized light as compared to disk. However, it allows the polarization plane position to be detected by the receptor. This is in line with the statement of Laughlin et al. (1975) that the orientation of dipoles along the microvilli is necessary mainly to obtain maximal absolute sensitivity to unpolarized light in rhabdomeric photoreceptors, and that their polarized light sensitivity appears to be no more than an extremely useful by-product of the evolution of absolute sensitivity. Fourth, when the disk membrane is rolled up to form a microvillus (see Moody and Parriss, 1961; Laughlin et al., 1975; Gribakin and Govardovskii, 1975), the latter is only 1.13 times more sensitive than solution, and only 0.75 as effective as disk for unpolarized light; even for linearly polarized light, with sensitivity of a microvillus being maximum, this system cannot be more sensitive than the disk (see Table I). According to Dartnall (1972), of many colored substances only astacene has a amax greater than that of

141

CELLULAR MECHANISMS OF INSECT PHOTORECEflION

cm2 versus 4.68 x cm", and this implies that the rhodopsin (9.9 X in both pigments approaches unity probability term of light absorption at A, which is the theoretical limit. Thus rhodopsin seems to be one of the most intensive stains known in organic chemistry. In conclusion, it can be said that the main factor contributing to the optical density of the photoreceptor (hence to its absolute sensitivity), namely, the molar TABLE I THEORETICAL DEPENDENCE OF AVERAGE MOLAREXTINCTION OF RHODOPSIN ON ORIENTATION OF ABSORBING DIPOLESFOR POLARIZEDAND UNPOLARIZED LIGHT" Average molar extinction at peak absorption, proportional to I,c,

+ luc, + Izcx

~~~~~~~~~~~

Microvillus model Mode of orientation of dipoles Components of concentration (Fig. 1) Polarized light I , = I , (or zero); I, = I, = 0 (or I, = l o ) Extinction for polarized light (literskm-mole) Unpolarized light I, = I, = I J 2 ; I, = 0 Extinction for unpolarized light (literdcm-mole) Maximal gain in extinction as compared to solution Polarized light Unpolarized light

Solution model c z = c, = C L = c/3

'h

IOC

Disk model c, = 0

c,

= cz = c/2

'h I,c

40,600

60,900

$5 I &

'h I,c

40,600

60.900

Made of disk membrane c, = c/4 cy = c/2 c, = c/4 'h I o c or % I,c

60,900or 30,450

Ideal (all-parallel)

c, = 0 cy = c c, = 0

I o c or zero

121,800,or zero %? I O C

45.675

60.900

1 .o

1.5

1.5

3.0

1.o

1.5

1.13

1.5

"The total c of pigment mobcules in the unit volume (see Fig. I) can be oriented either randomly in the volume (solution model) or randomly in the yOz plane (disk model) or all parallel to the y-axis (ideal microvillus). The intensity I , in the case of unpolarized light can be considered as consisting of two components I, = I, = 1,/2 with mutually perpendicular E vectors. In the case of linearly polarized light either the I, or I, component is equal to the full intensity I o, whereas the other two are zero. Numerical values for molar extinction are from the basic value determined for a solution as 40,600literslcm-mole by Wald and Brown (1953).

142

F. G. GRIBAKIN

extinction E (A, (o) of rhodopsin, has a very high value. Absolute sensitivity of the rhabdomeric eye strongly depends on the orientation of the rhodopsin molecules in the photoreceptor membrane and, in the case of ideal alignment of the molecules along the axes of the microvilli, the value of the average molar extinction (and the same is true for the molecular extinction coefficient) is equal to that of vertebrate rod and cone disks for unpolarized light. Also, sensitivity to polarized light arises which can be considered a by-product of evolutionary growth in absolute sensitivity of the compound eye photoreceptors (Laughlin et al. 1975). In the above discussion the molar extinction of insect rhodopsin was considered the same as that in vertebrates and cephalopods (see Goldsmith, 1972). b. Concentration of Rhodopsin. The concentration of rhodopsin in vertebrate photoreceptors is known to be 2-2.5 mM (see Liebman, 1972), whereas for an arthropod Hays and Goldsmith (1969), taking the molar extinction to be 40,000, obtained 1.4 mM. Specific absorbances (per unit length of rhabdomere) at,,,A appeared to be nearly the same in different arthropod species (with all the values determined for the E vector parallel to microvilli): 0.0073 in the crab Libinia (Hays and Goldsmith, 1969), about 0.0085 in the crayfish Orconectes (calculated from the data of Waterman et a/., 1969), and 0.0075 in the fly (Kirschfeld, 1969). From this similarity one can assume that the rhodopsin concentration in the photoreceptors of different arthropods must be nearly equal (about 1.5 mM). In favor of this assumption is a similar density of protein particles in the photoreceptor membranes of different arthropods (7000/pm2; see Section I1,C). The upper limit of rhodopsin concentration in the rhabdomeric membrane can easily be determined. Let the unit volume (1 dm3 or liter) be completely filled with spherical rhodopsin molecules 8 nm in diameter (like protein particles in a microvillar membrane) forming a hexagonal or cubic lattice. Then the molar concentration of these molecules is c = k (VO/VIn) (1/N)

(6)

where c is the molar concentration, V ois the unit volume (1 liter = lP4nm3),Vm is the volume of one molecule (2.7 x 10' nm3), N is Avogadro's number, and k is the relative volume occupied by spherical molecules within the unit volume. The value of k can easily be calculated and is rr *I6 = 0.74 for a hexagonal lattice and T I6 = 0.52 for a cubic lattice. The calculated rhodopsin concentrations given by Eq. (6) are 4.56 and 3.25 mM for hexagonal and cubic lattices, respectively. However, in a rhabdomere consisting of microvilli 50 nm in diameter only 0.48 of the volume is taken up by the photoreceptor membrane (Gribakin and Govardovskii, 1975). Then the upper limit for the rhodopsin concentrations in the rhabdomere must be reduced to 2.18 and 1.56 mM, respectively. The second value appears to be very close to the above microspectrophotometricdata. Thus the second factor contributing to the optical density D, namely, the rhodopsin concentration c, is about 1.5 mM in arthropods, and this value is only

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

143

30% less than that characteristic of the most dense hexagonal lattice and conforms rather to a square lattice. c. The Length of the Rhabdomere. The length of the rhabdomeres in arthropods may range from several micrometers to several hundred micrometers; for instance, it is 400 p m in the crab Callinectes (Eguchi and Waterman, 1966), and 400 pm in the drone bee (Perrelet, 1970). With a specific absorbance of 0.0078/pm (Hays and Goldsmith, 1969) and a length of 300 pm, the transmission of the rhabdomere is 4.5 x l o + , or less than 0.5%, which means a relative absorption of more than 0.995; thus the length of the rhabdomere in arthropods seems to be the most changeable factor of the three related to optical density, and consequently to the absolute sensitivity of the photoreceptor. In order to provide high absolute sensitivity of the photoreceptor, a rhabdomere can evolve to become long enough so that it can absorb every quantum entering its distal end with a probability approaching unity. 3. Waveguide Effects and Sensitivity Control In Section II,D,2 no special mention was made of a mechanism which could confine a light wave, once it enters a thin absorbing fiber (the rhabdomere), to within this fiber, thus causing it to travel the full fiber length. Dielectric waveguide theory, the fundamentals of which were elaborated in the 1960s as the basis of fiber optics technology and exploration, provides a clue to this mechanism. As applied to photoreceptors of insects and other arthropods, the theory was advanced by A. W. Snyder and his co-workers in about 30 papers published during the last 10 years. The general principles of waveguide optics as applied to absolute light sensitivity of the rhabdomeric photoreceptor are considered briefly (for full theory, see Snyder, 1975a,b) in the following discussion. a. General Ideas of Waveguide Optics. Any optical fiber made of a material with a refractive index n1 and immersed in or surrounded by a medium with a refractive index n2 < n , may serve as a light guide if illuminated end on. When transmitted along the fiber, light, being an electromagnetic wave, forms specific field patterns termed dielectric waveguide modes. Every mode is a consequence of resonant or interference effects derived from internal reflections of the light wave within the fiber and can be characterized by the field distribution in a cross section of the fiber (Snitzer, 1961; Enoch, 1963; Snyder, 1975b). Only a fraction of the light energy is transmitted by a given mode within the fiber; the rest is transmitted outside the fiber, though along it, and fails to contribute to absorption in the fiber. The most important parameter of the fiber determining the number of modes and the fraction of energy transmitted within it is the so-called characteristic waveguide parameter I/: where d is the diameter of the fiber, h is the wavelength, and n , and n2 are the refractive indexes of the fiber and medium, respectively. A limiting value of V

144

F. G. GRIBAKIN

FIG.3. Relative power P channeled through the optic fiber versus the waveguide characteristic parameter V for the angle of incidence 0 = 0. Replotted from the data of Snyder (1975b).

exists for each mode (the so-called cut-off V or V c ) , and at V < V , the given mode fails to propagate. Snyder (1975b, see his Fig. 9.8) has calculated a fraction of the total light power P(V,8) transmitted by all the modes within a fiber versus V and the angle of incidence 8. From his data, taking 8 = 0 (an axial light beam), we can plot P(V,t?) as shown in Fig. 3 and then evaluate the fraction of light channeled through real rhabdomeres and accessible to interact with rhodopsin. b. Waveguide Properties of the Rhabdomere. The refractive indexes of photoreceptor structures lie between those of the bilipid layer (1.66) and of Ringer’s solution (1.336) (Table 11). In arthropods the highest refractive index for a rhabdomere has been reported to be 1.405, and the lowest 1.347 (Table 11). From these extreme values the characteristic waveguide parameter V is calculated as given in Table I11 for h = 500 and 350 nm. Since for V 2 4 more than 80% of the light energy is transmitted within the rhabdomere (Fig. 3), we can infer that, first, the waveguide properties of the rhabdomere do not reduce markedly the absolute senstivity of a photoreceptor at V 2 4 and, second, since they provide confinement of light to within the rhabdomere, the Beer-Lambert absorption law remains applicable for the rhabdomere, taking its length as the absorbing layer thickness (see Section II,D,2). Thus the optical properties of the rhabdomere allow it to function as a photon counter at low light intensities, and the problem arises of how to lower the absolute sensitivity of the photoreceptor at high intensities. 4. Optical Mechanisms of Absolute Sensitivity Control

The effects of light adaptation on the microenvironment of the rhabdomere are well known, and they are, first, dispersion of the principal endoplasmic cistern (PEC) into the peripheral cytoplasm (“palisade” movement) with mitochondria migrating to the rhabdom and, second, radial migration of screening pigment

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

145

granules to the rhabdom (for review, see Walcott, 1975). The former results in a refraction change in the medium surrounding the rhabdom (or rhabdomere), while the latter modifies both the refraction and, mainly, absorption properties of the rhabdom “cladding.” Both mechanisms are intended to reduce the fraction of light absorbed in the rhabdomere. a. Refraction Waveguide Mechanism of Sensitivity Control. As seen in Fig. 3, high absolute sensitivity of a photoreceptor can easily be reduced by a decrease in V, which means that a fraction of incident light power is allowed to leave the rhabdomere or even pass by it. Perhaps during palisade migration (porridge and Barnard, 1965), of all the terms contributing to V, only n2 changes; this has two consequences, namely, a change in V and in the acceptance angle. Since for efficient n,-dependent sensitivity control V must be less than 4 (see Fig. 3), we can express this condition, from Eq. (7), as

V which can be rewritten as

=

(.rrd/A)(n12- n22)1’25 4

(8)

(n2/n1)22 1 - (4A/r dn,)2

(9) This means that, the greater the diameter of the rhabdom (i.e., the greater the dlX ratio), the less the difference between n2 and n, must be to provide sensitivity TABLE II REFRACTIVE INDEXES OF PHOTORECEFTOR STRUCTURES ~~

Refractive index Bilipid layer Ringer’s solution Rod outer segment Rod outer segment photoreceptor membrane Rod outer segment cytoplasm Bee rhabdomere Fly rhabdomere Bee cytoplasm Fly cytoplasm Surrounding media Bee PEC Fly axial cavity Effective surroundings in fly (cavity plus cytoplasm)

1.66 1.336 1.41

Reference‘‘ a b C

1.475 1.365 1.347 1.349 1.365 1.390-1.405 1.343 1.340 1.339 1.336

d e

1.339

f

“(a) Finean and Engstrom (1967); (b) Liebman er al. (1974); (c) Sidman (1957); (d) Varela and Wiitanen (1970); (e) Seitz (1968); (0 Stavenga (1974); (g) Kirschfeld and Snyder (1975).

146

F. G. GRIBAKIN

TABLE III WAVEGUIDE CHARACTERISTIC PARAMETER V CALCULATED FOR SEVERAL ARTHROPOD SPECIES" Initial parameters Waveguide (rhabdomere) diameter (pm) n , = 1.405; n 2 = 1.339 For A = 500 nm For A = 350 nm n = 1.347; n 2 = 1.339 For A = 500 nm For A = 350 nrn

,

~~~~~~~~~~~~~~~~~~~~

Bee

fly

Cricket

Crayfish

1

2

2

4

15

2.54 3.64

5.08 7.28

5.08 7.28

10.16 14.56

38 55

0.91 1.30

1.81 2.59

1.81 2.59

3.62 5.18 ~~~~~~~

13.5 19.4 ~~

"The highest and lowest n , - n 2 differences are from Table 11. Diameters of rhabdomeres (rhabdoms) are from Trujillo-Cenoz and Melamed (1966) for the fly, Gribakin (1967) for the bee, Polyanovskii (1976) for the cricket Gryffus, and Eguchi and Waterman (1966) for the crayfish Orconectes.

control through a change in the refraction of the surroundings. For instance, for n , = 1.4 and d = 10 pm, V can be made less than 4 only if n2/nl = 0.9990 (a 0.1% difference in refraction), whereas for d = 2 pm, n 2 / n 1must be as large as 0.9900 (with a 1% difference between n2 and nl). From Table I1 one can see that the real difference between n , and n2 is about 1%, hence sensitivity control through a change in the refraction of the surroundings works efficiently only in rhabdoms no more than 3-4 p m in diameter. For instance, consider the bee rhabdom (see Table 11) with nl = 1.347 and a diameter of 2 p m (for wavelength 500 nm). Substitution of the dark-adapted refractive index n2 = 1.339 (bee endoplasmic cistern) for n2 = 1.343 (bee cytoplasm) in the light causes a decrease in V from 1.81 to 1.28 and consequently a change in the relative amount of light power absorbed from 0.44 to 0 . l b n e a r l y a three-fold decrease in sensitivity. (Note that nl = 1.347 is only 1% more than n = 1.336 for Ringer's solution). This mechanism probably provides sensitivity control over a wider range when organelles having a refractive index higher than that of cytoplasm (e.g., mitochondria) surround the rhabdom in the light, and these structural changes have been found in the light-adapted photoreceptors of locusts (Homdge and Barnard, 1965) and crickets (Petrosyan, 1977a). Simultaneously with a decrease in the fraction of light power confined to the rhabdom, the acceptance angle is changed. As found experimentally, the acceptance angle of the photopic eye is reduced approximately from 6"-7" in the dark to 2.5"-3.5" in the light (Tunstall and Honidge, 1967, locust; Butler and Hor-

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

147

ridge, 1973, cockroach). This should bring about attenuation of the relative light power within the rhabdom nearly proportional to the sine squared of the acceptance angle, or approximately 5 to 10 times to that in the dark. Theoretical analysis of angular sensitivity can be found in Snyder (1975b). Thus the refraction mechanism of absolute sensitivity control might lower the absolute sensitivity by at least one order of magnitude, or even more,, and this seems to be of importance in insects with no radial pigment migration (e.g., the locust). The refractive index of the bee rhabdom (1.347) may appear very low in comparison with that of the fly (formerly 1.340 which has been corrected to 1.365 and even to 1.405; see Table 11). Nevertheless, from these values the solid content of rhabdomeres can be determined: c, = [(nl - n , ) / a ] x 100

(10)

where c , is the solid content in percent, n, and n , are the refractive indexes of the rhabdomere and of water, and a is the refractive index increment (0.18 for biological tissues; see Finean and Engstrom, 1967). Thus the solid content of the bee rhabdom is as low as 6.1%, while those for fly rhabdomeres and rod outer segments are similar-38 and 41%, respectively. Further, for the bee rhabdom, assuming a rhodopsin concentration of c = 1.5 mM and a molecular weight as low as M = 50,000 daltons (Section LII,D), we obtain the solid content: c, = (Mc/IooO) x

loo = 7.5%

(11)

which means that the refractive index for the bee rhabdom is likely to require correction, since the dry weight of only rhodopsin (7.5%) exceeds the total solid content of the rhabdom found with refractometry (6.1%) performed by Varela and Wiitanen (1970). Nevertheless, the refractive mechanism of sensitivity control seems to work successfully in insects and other arthropods, however the refractive indexes of their rhabdomeres are defined. b. Absorption Waveguide Mechanism of Sensitivity Control. This mechanism works when screening pigment granules migrate to the rhabdom in the light, hence both the refraction and absorption of optical fiber caldding are changed. According to the analysis of Snyder and Homdge (1972) and Snyder (1975b) as applied to the cockroach rhabdom, only 25% of a 10-fold decrease in sensitivity in the light is due to the refraction mechanism, while the rest is produced by the absorption of light transmitted outside the rhabdom in pigment granules. The absorption waveguide mechanism of sensitivity control was first described in terms of photoreceptor optics by Kirschfeld and Franceschini (1969) in Musca. In this insect it takes only a few seconds for the sensitivity to decrease and pigment granules to congregate close to the rhabdomeres in cells Nos. 1-6 after the light is switched on. Recently, Stavenga and Kuiper (1977) and Stavenga et al. (1977) showed experimentally that the same mechanism operates in hymenopterans and in butterflies where it covers about 3 log units of sen-

148

F. G. GIUBAKIN

sitivity with a time constant of only a few seconds as in the fly. The time constant in hymenopterans appears to be about 10 seconds. The absorption waveguide mechanism of sensitivity control has been termed as “insect pupil mechanism” by Stavenga and co-workers, since in the light screening pigment forms a kind of “longitudinal pupil” around the rhabdom. Screening pigment material of cockroach photoreceptors has been calculated to be 70 times more absorptive than the material of the rhabdomere (Snyder, 1975b) though, as one can judge from the data of Langer (1975), the specific zbsorbance of single primary pigment cell granules is lower-about 0.0275/pm in Gerris and 0.1000/pm in Apis, which is, respectively, 3.5 and 13 times greater than that of the rhabdomere (about 0.0075/pm; see Section 111,D). Unfortunately, data on the specific absorbance of individual pigment granules of photoreceptors are not available because of their small dimensions. Thus two waveguide mechanisms provide a means for absolute sensitivity control and, since these manifest themselves as ultrastructural changes in the perirhabdomeric region of the photoreceptors, it is reasonable to consider both mechanisms in terms of cell biology. 5. Cell Biology and Sensitivity Control a. Cellular Manifestations of Sensitivity Control. In general, cellular manifestations of absolute sensitivity control (or dark-light adaptation) is not confined to migration of the palisade, mitochondria, and pigment granules in photoreceptors themselves-the effects most characteristic of the photopic compound eye. Other known structural adaptations are longitudinal migration of screening pigment in photoreceptors and pigment cells (both primary and secondary), movement of photoreceptor cell bodies and their rhabdomeres, and movement of cone cells and their crystalline tracts (see, for review, Horridge, 1975; Walcott, 1975). The time necessary for the above processes to be accomplished after the light is switched on are quite different: from a few seconds (fly, Kirschfeld and Franceschini, 1969) to about a minute (wasp and bumblebee, Stavenga and Kuiper, 1977) for radial pigment migration in photoreceptors; 10-15 minutes for palisade movement (locust, Horridge and Barnard, 1965; Tunstall and Homdge, 1967); 10-30 minutes for longitudinal pigment migration (moth, Post and Goldsmith, 1965); and 40-60 minutes for cone and photoreceptor cell movement (Eckert, 1968; Walcott, 1971). Apart from more-or-less detailed descriptions, nothing is known at present about the nature of these photomechanical mechanisms of sensitivity control, and only the initial steps have been made in understanding probably the simplest mechanism-light-evoked migration of pigment granules and palisade movement (see Miller, 1975). b. Light-Dependent Pigment Migration. When illuminated, pigment granules in both photoreceptors and pigment cells change their positions; they tend to congregate close to the rhabdom in photoreceptors of the photopic eye and move proximally from a distal dark position to form a continuous pigment

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

149

sheath around the retinula in the secondary pigment cells of the scotopic eye. The general features of pigment migration in both the photopic and scotopic eye appear to be similar, and they are as follows (see, for review, Goldsmith and Bernard, 1974; also Bernhard and Ottoson, 1964; Post and Goldsmith, 1965; Hoglund, 1966; Kirschfeld and Franceschini, 1969; Butler, 1971; Menzel and Lange, 1971; Franceschini, 1972; Kolb and Autrum, 1972, 1974; Menzel and Knauth, 1973, etc.). The movement of pigment is restricted to the illuminated ommatidia, or even to individual illuminated cells within the same ommatidium (including cells selectively adapted with monochromatic light), and this opposes any humoral or neuronal control of pigment migration. Carbon dioxide and narcosis make pigment granules move to light-adapted positions in both photopic and scotopic eyes, and so does cold in scotopic eyes. The extent of pigment displacement to the light-adapted position depends on the intensity of the light. Using action spectra of pigment movement dependent on wavelength and polarization, Franceschini (1972) has stated that light-induced pigment migration in the photoreceptors of the photopic eye is triggered by visual pigment itself, but little is known about the nature of the mechanism of motion. The light-induced depolarization of the photoreceptor is believed to evoke pigment migration, and consequently two hypotheses have been put forward to explain the forces which could drive pigment granules in the cell (Kirschfeld and Franceschini, 1969). According to the first (the passive movement hypothesis), granules are forced to move to the rhabdom by an electric field dependent on light-induced depolarization of the cell (see also Stavenga, 1971; Walcott, 1975). An alternative mechanism (the active movement hypothesis) is thought to be connected with the activity of microtubules (see also Miller, 1975) in a manner similar to that observed, for instance, in fish melanophores (e.g., Bikle et al., 1966). While there is no evidence to support the first view as yet, Miller and Cowthon (1974) and Miller (1975) stimulated light-induced palisade movement and radial pigment migration in photoreceptors of Limulus using colchicine, and a simultaneous decrease in the number of radially orientated microtubules was observed. However, treatment with colchicine did not mimic other anatomical changes generally evoked by light in Lirnulus photoreceptors, namely, an increase in the number of small multivesicular bodies and a decrease in the thickness of the rhabdomeres. Colchicine, being an antimitotic agent, exerts disruptive effects on microtubules in vivo (e.g., Wikswo and Novales, 1972), and its lightlike action upon pigment movement might mean that the microtubular system of the photoreceptor, when disrupted by colchicine, cannot confine pigment granules to the periphery of the cell any longer as it can in the dark. A mechanism triggering pigment movement in the secondary pigment cells of the scotopic eye has not been described, and many arbitrary possibilities can be offered. For instance, movement could be due to absorption of light by the

150

F. G. GRIBAKIN

pigment granules themselves, a change in the ionic composition of the extracellular space between the photoreceptor and the pigment cell which might modify the resting potential of the pigment cell, the release of a chemical transmitter or messenger from the photoreceptors, and so on. It is interesting, however, that in the dark all the pigment granules congregate to form a dense mass in the distal region of the cell that is in its perinuclear region. Similarly, in the dark, fish melanophores show “contraction,” and their pigment also is confined to the perinuclear region of the cell (Bikle ef al., 1966). In connection with this it is useful to recall briefly what is known about pigment migration in fish melanophores (Bikle et al., 1966; Egner, 1971). In isolated scales from the dorsal side of Fundulus heteroclitus 0.1 M potassium chloride and adrenaline ( 10-3-10-5 M) cause “contraction” of melanophores, whereas 0.1 M sodium chloride causes “expansion. ” Usually, granules move along more-or-less limited paths in the cytoplasm of the melanophore processes. It seems that granules on parallel paths move nearly independently, while those. on the same path can move as single units (chains) consisting of three to five granules. Occasionally, during a short period of time units on neighboring paths may move in opposite directions. Electron microscopy shows that, in the processes of melanophores, granules are lined up along microtubules, and this is probably due to the motive function of the latter. Microtubules diverge radially from a certain locus in the cell body where a centriole is usually located, though direct connection between microtubules and the centriole has not been found. The activity of melanophores is controlled by the nervous system, and the membrane potential does not seem to be a factor causing granules to move (Egner, 1971). With reference to secondary pigment cells, an assumption should be made that the dark-adapted position of the pigment imposes a greater requirement upon the metabolic energy of the cell (cf. carbon dioxide, narcosis, and light action; see also Goldsmith and Bernard, 1974). Whether microtubules control pigment movement in these cells is still unknown. c. Palisade Movement. The term “palisade,” introduced by Horridge and Barnard (1965), is not exact and from a cytological point of view this formation is the principal endoplasmic cistern (PEC) of the photoreceptor (Gribakin, 1975). However, workers in the field often call the PEC a palisade in the literature. From the refractive index of the PEC (1.339 in the bee; see Table 11) one can infer its solid content to be about 1.7%,while that of the cytoplasm is 4% (n = 1.343). Also, the potassium content of the PEC is lower than that of the cytoplasm (Gribakin et al., 1976; Burovina et al., 1978). Thus the PEC content is mainly water, and PEC contraction and dispersion into smaller cisterns in the light, termed palisade movement, is probably accompanied by water transport. It is interesting that other processes similar to pigment and palisade movement are believed to be connected with the transport of water in different tissues from Calliphora salivary glands to the toad bladder (Berridge and Prince, 1972; Taylor

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

151

et al., 1973). These processes are also related to the action of microtubules and microfilaments which on being inhibited by antimitotic agents (colchicine, vinblastine, podophyllotoxin) cease responding to the antidiuretic hormone vasopressin (Taylor et al., 1973). The fact that in vivo transport of water is also cyclic-AMP- and calcium-dependent suggests that the mechanism of light action on both pigment migration and palisade movement has a striking resemblance to the action of hormones. The existence of a light-dependent extracellular palisade, discovered by Meyer-Rochow (1974) in the compound eye of the beetle Creophilus erythrocephalus, makes this idea even more intriguing. d. Energetics of Sensitivity Control and Entropy of the Compound Eye Structure. The above discussion of the cellular manifestations of sensitivity control closely related to optical mechanisms shows that the compound eye, when darkadapted, displays more ordered fine structure (lower entropy) than when lightadapted. Indeed, in the course of adaptation to the dark, the photopic eye photoreceptor seems to pump liquid (water?) into its perirhabdomeric cistern, and the PEC thus formed (like a balloon) presses pigment granules andor mitochondria back from the rhabdom. This process should require metabolic energy supplied by the cell. In the light, the PEC liquid become free to disperse into the cytoplasm as small cisterns, and pigment andor mitochondria are allowed to approach the rhabdom. Similarly, during dark adaptation the pigment cell of a scotopic eye retracts all the pigment granules back into its distal region where a dense pigment cluster forms, and again this process should show an energy demand, since the resulting distribution is far from equlibrium. In the light, pigment granules become more fkee to drift toward the proximal end of the cell, the distribution approaching uniformity and the process resembling diffusion. Thus, considering the change in morphology alone, one could already come to the important conclusion that, in order to realize the high absolute sensitivity inherent in the light-absorbing structures of the photoreceptor, an insect must continuously supply the eye with metabolic energy, thus providing perfect structural, hence optical, conditions for absorption. In other words, energy consumption in the dark should be more than that in the light. Two experimental findings support this view. First, as Autrum and Tschamtke (1962) have found, oxygen consumption in the dark is higher than in the light in the locust and cockroach compound eye, that is, in insects showing well-expressed movement of the palisade, pigment, and mitochondria. This is also true for the bee eye, but only at temperatures not exceeding 25°C. The opposite has been found for the Calliphoru eye (see also Hamdorf and Schwemer, 1975), which has no palisade movement but only radial pigment migration, and for the bee eye at 34°C. Second, carbon dioxide, narcosis, and cold make the screening pigment pass into the light-adapted position, and this is likely due to a deficit in metabolic energy (cf. Goldsmith and Bernard, 1974).

152

F. G. GRIBAKIN

Thus we can infer that, when dark-adapted, the compound eye exhibits the lowest structural entropy and an energy consumption higher than when lightadapted, and in connection with this a conclusion may be given: the adaptation of the insect eye to the dark is the process requiring the continuous metabolic energy influx, which is necessary for the formation and preservation of perfect optical structures for a prolonged period of time; the adaptation of the eye to the light is the light-protective response to the excessive stimulation of the photoreceptors, whose normal function is photon counting. The light-protective mechanisms are on automatically, and no additional energy seems to be required. This conclusion emphasizes the principal difference between two main photobiological processes-photosynthesis and vision. Using the former, the organism utilizes the light energy. By contrast, the latter requires the energy from the organism (cf. Section II,C,2,c and Vinnikov, 1974). e. Light-Evoked Structural Changes Connected with Rhabdomeres. There are many light-evoked structural changes in rhabdomeric photoreceptors which seem to have a lesser beating on active sensitivity control; they are swelling of rhabdomeric microvilli (Rohlich and Torok, 1962; Rohlich and Toro, 1965; Gribakin, 1969a,b, 1975), diminution of the rhabdomere (White, 1967, 1968; White and Lord, 1975; Miller and Cowthon, 1974), and an increase in number of cytoplasmic organelles such as multivesicular bodies, lamellar bodies, and mixed vesicular-lamellar bodies (Eiguchi and Waterman, 1967, 1968; White 1967, 1968; Eguchi et al., 1973; Miller and Cowthon, 1974; Miller, 1975; Nemanic, 1975; Behrens and Krebs, 1976). All these changes are thought to be connected with destruction of the photoreceptor membrane in the light and its renewal during dark adaptation (Burnel et al., 1970). Destruction of the photoreceptor membrane in arthropods is believed to occur by phagocytosis at the bases of the rhabdomeric microvilli. The mechanism of photoreceptor membrane renewal is still totally obscure (Behrens and Krebs, 1976). The above changes in structure are usually obtained using high light intensities andlor prolonged illumination. For this reason, they could hardly be related to mechanisms which control absolute sensitivity in the natural light environment of the given species. Nevertheless, ultrastructural changes in rhabdomeres produced by intense and/or prolonged illumination proved to be of much use, since they have made it possible to discover cellular bases for color vision and polarized light perception in arthropods (J3guchi and Waterman, 1968; Gribakin, 1969a,b, 1975; Eguchi et al., 1973). 6. Conclusion The compound eye intended to operate in the light environment usual for given species has evolved rhabdomeres which can normally work as photon counters. To attenuate a light flux reaching its rhabdomeres from a brighter environment, the eye triggers cellular mechanisms, which optical and electrical functions are

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

153

to decrease its absolute sensitivity so that the amount of light which interacts with visual pigment within the rhabdomeres can be reduced to a level close (but not identical) to the original one. As quantitative comparison has shown (Kirschfeld, 1974), an insect rhabdom receives nearly the same (or even a greater) number of quanta as a vertebrate photoreceptor when operating in the same light surroundings, however, the compound eye pays for high absolute sensitivity, combined with extremely small size, with a loss in angular resolution (Kirschfeld, 1974).

E. COLORVISION 1. General Approach

It seems evident that the great variety of insect visual pigments considered in Section II,C,2 is a prerequisite for color vision in these animals. Moreover, since all insect visual pigments include retinal as a chromophoric group, their absorption spectra have to follow Dartnall’s nomogram (Dartnall, 1953). However, photoreceptors incorporating Dartnall’s visual pigments usually show spectral properties different from those of their native pigment, and this difference is due totally to geometry (or, which is the same, morphology) of both the rhabdom and its rhabdomeres. Since the theoretical principles of these effects have been elaborated rather well (Snyder, 1975a,b; Bernard, 1975; Gribakin and Govardovskii, 1975), only a brief account of the cellular mechanisms of color vision in insects is given here.

2. Isolated Rhabdomeres and Self Screening of Visual Pigmen2 An isolated rhabdomere is usually referred to as a “fly rhabdomere” (Snyder, 1975a,b; Gribakin and Govardovskii, 1975), and this is considered an isolated absorbing optic fiber. It is obvious from Eq. (I) (Section II,D,2) that the shape of the absorption spectrum of an isolated rhabdomere strongly depends not only on the pigment extinction coefficient a ( X , cp) but also on the product c x I (concentration times rhabdomere length). In the limit, at c X I -+ 03, all the light in the pigment absorption band could be absorbed by the rhabdomere, and consequently the absorption spectrum of the rhabdomere would show no dependence on wavelength (so-called self-screening). In other words, the greater is c X I , the greater the absolute sensitivity, but the less the spectral selectivity of the photoreceptor (see Snyder, 1975a,b; Gribakin and Govardovskii, 1975). This statement is quantitatively illustrated in Fig. 4 and 5 where real values for the extinction coefficient and pigment concentration have been taken from the results in Section I1,D. Thus the above discussion indicates that the perfect spectral selectivity necessary to evolve color vision is in conflict with the high absolute

154

F. G. GRIBAKIN

400

500

A . nm

600

FIG.4. Relative absorption A of an isolated rhabdomere versus wavelength A calculated for different rhabdomere lengths-50, 100, 200, and 300 p n . The molar extinction of rhodopsin at A ,,,ax = 500 nm has been taken equal to 40,600 litedcm-mole (Table I), and the rhodopsin concentration as M (from Section III,D,2). Waveguide effects have been ignored. 1.5 x

sensitivity requirement. In many insects this situation has been mastered by the evolution of a fused rhabdom. 3 . Fused Rhabdoms, Lateral Filtration, and Color Vision

A fused rhabdom consisting of rhabdomeres with the same visual pigments has no advantage in spectral selectivity, and from this viewpoint is totally identical to an isolated rhabdomere. This is not true if the rhabdomeres comprising the fused rhabdom contain different rhodopsins. The first experiment demonstrating different spectral cell types in the ommatidium of the honeybee worker was that of Gribakin (1969a,b). The same appeared to be true for other insects (Mote and

300

400

500

A . nm

600

FIG. 5. The relative absorption spectra A in Fig. 4 normalized to their maxima at 500 nm to illustrate the deterioration of spectral selectivity with increase in rhabdomere length (respectively, 50, 100, 200, and 300 pm). Rh is the relative absorption of rhodopsin.

CELLULAR MECHANISMS OF INSECT PHOTORECERION

155

Goldsmith, 1971; Butler, 1971, for the cockroach; Menzel and Knauth, 1973, for the ant; Kolb and Autrum, 1974, for the bee). The first theory concerning the fused rhabdom was elaborated by Snyder el al. (1973) who showed that the presence of more than one spectral type of rhabdomere within a fused rhabdom narrowed the spectral sensitivity curve of a photoreceptor because neighboring rhabdomeres acted as colored light filters for each other. Since every filter extends the full length of the rhabdom, the effect has been termed lateral filtration (Snyder et al., 1973). Further details can easily be found in the original paper, and we strongly emphasize the fact that the fused rhabdom formed by rhabdomeres with different visual pigments proved to be one of the most efficient developments in the evolution of the insect visual system which allowed perfect color vision to evolve without any loss of absolute sensitivity (Snyder et al., 1973). 4. Tiered Rhabdoms

Tiered rhabdoms are known to exist in many insects, but their physiology is not clear yet, nor is their optics. With respect to color vision, longitudinal filtration may take place in tiered rhabdoms with effects somewhat similar to those of lateral filtration. If one tried to calculate the relative absorption curves for a tiered rhabdom consisting of rhabdomeres with the same visual pigment, one would obtain curves like those shown in Fig. 6, which have never been seen experimentally. When investigating spectral sensitivity curves for the cricket Gryllus domesricus, we failed to find different color receptors and, since the animal has a typically tiered rhabdom (Polyanovskii, 1976), one would expect to obtain curves like those in Fig. 6. However, we have never seen them either (F.

h , nm

FIG.6 . Calculated normalized relative absorption spectra of proximal rhabdomeres, showing the probable effect of longitudinal filtration in a tiered rhabdom. The proximal rhabdomere is taken to be 100 k m in length, while the distal one is 50 or 1 0 0 pm long. Both rhabdomeres contain the same rhodopsin.

156

F. G. GRIBAKIN

G. Gribakin and T. M. Vishnevskaya, unpublished). Perhaps the two-peaked receptors with both peaks in the visible region (Eguchi, 1971, tiered rhabdom of a dragonfly Aeschna) can be explained in this way (Gribakin and Govardovskii, 1975) but, as measured by Eguchi, the peaks proved to be much sharper than the theory predicted, and consequently further study along these lines is needed. An alternative explanation of double-peaked sensitivity curves having no relation to the tiered rhabdom is off-axis illumination (Snyder, 1975b).

5 . Comblike (‘ ‘Crustacean-Like”)Rhabdoms Comblike rhabdoms are not typical of insects, though they have been found in several species (Meyer-Rochow, 1974, staphylinid beetle Creophilus; Frantsevich et al., 1977, the lamellicornian beetle Lethrus). Nothing is known about spectral cell types of the Creophilus retinula, while Lethrus has been found to have color vision (Frantsevich e f al., 1976, 1977) which is mediated by longwave and ultraviolet receptors. Unfortunately, we failed to observe structural changes in the Lethrus retinula during selective color adaptation (A. D. Polyanovskii and F. G. Gribakin, unpublished), and this is why the interdigitation of microvillar layers in the Lethrus rhabdom cannot be explained at present in terms of color vision. In crustaceans, however, both longwave and shortwave receptors contribute to both the sets of mutually perpendicular rhabdomeric microvilli (Eguchi et al., 1973). Since in the crayfish retinula color receptors of the same modality occupy no more than two quadrants of the square rhabdom cross section, one can assume that lateral filtration takes place in comblike rhabdoms as well. Thus we seemingly have another example of a photoreceptor system displaying color vision combined with high absolute sensitivity.

6. Double-Peaked Sensitivity Problem Wasserman (1973) has paid special attention to broad-banded double-peaked photoreceptors with the peaks located usually in the visible and ultraviolet regions, or in the green and blue regions (beta cells, unlike “tuned” alpha cells). Beta cells do not seem to be due to artifactual recordings from two different cells simultaneously, and three of many explanations may be valid: (1) self-screening of visual pigment (Section II,E,2 and Wasserman, 19731, (2) electrical coupling (Snyder et al., 1973), and (3) sensitising pigments’ (Kirschfeld et al., 1977). Broad-banded beta cells might give an advantage in a poor light environment when every quantum absorbed, no matter what its energy (wavelength), provides information on the presence of a light source. Comparative analysis of light quanta by their energies, that is, color vision, has no meaning under these conditions because of the shortage of light quanta to be compared (cf. scotopic and photopic vision in humans). Indeed, we think that the much more serious problem in color vision is to obtain narrow-banded receptors, since every visual pigment, apart from its a-absorption peak in the visible region, must display a

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

157

P-absorption peak in ultraviolet light. Nothing is known about how this problem was solved early in evolution in both vertebrates and invertebrates. Gribakin and Govardovskii (1975) have suggested that rhodopsin molecules, known to have their a-and @absorbing dipoles nearly perpendicular, might be incorporated into the photoreceptor membrane so that their f3 dipoles can avoid interaction with light waves (e.g., lie in parallel with the light path). Perhaps still unknown filtering effects suppress p absorption, or insects have mastered the technology of making “tuned” visual pigments with no additional maxima (as they have in the case of ultraviolet-sensitive rhodopsin; see Section I1,C). 7. Conclusion As the above brief survey has shown, physiological characteristics of insect color vision systems are closely related to the morphological organization of the ommatidium and its photoreceptor membrane, and such complicated evolutionary developments as fused, comblike, and probably tiered rhabdoms serve to overcome contradictory requirements for color vision and absolute sensitivity.

F. SENSITIVITY TO POLARIZED LIGHT Polarization sensitivity (PS) in arthropods is thought to be a by-product of the high absolute sensitivity demand in the compound eye with rhabdomeric photoreceptors (Laughlin et al., 1975; see Section II,D) and is totally due to the orientation of rhodopsin molecules in the photoreceptor membrane or, in other words, to dichroic properties of the membrane. The theory of polarized light perception in arthropods has recently been presented in detail (Snyder, 1973, 1975a,b; Gribakin, 1973, 1975; Gribakin and Govardovskii, 1975; Waterman, 1975; Menzel, 1975a), and the reader is referred to the literature for the fundamentals of the theory. Here we give only a simplified outline of the problem. An individual rhabdomeric microvillus made of a universal photoreceptor membrane should show a dichroic ratio of 2 (Moody and Parriss, 1961). In the case of an isolated rhabdomere, the longer the rhabdomere, the lower its dichroic ratio, because of rhodopsin self-screening (Shaw, 1969b). Thus an isolated rhabdomere made of a disk membrane fails to display a PS of more than 2, and this is true for cells 1 through 6 of the dipteran retinula (Kirschfeld and Snyder, 1975). Waveguide effects may increase the PS of a thin, isolated rhabdomere, and this probably occurs in central dipteran rhabdomeres (R7 and R8) which are as small as 1 Fm in diameter (Snyder, 1973). A comblike (crustacean-like) rhabdom makes it possible to gain PS identical to the dichroic ratio of a single microvillus without any loss in absolute sensitivity (plus color vision) (Shaw, 1969b; Snyder, 1973). The fact that photoreceptors with comblike rhabdoms in insects mainly show a PS of about 2 (Meyer-

158

F. G. GRlBAKW

Rochow, 1974, Creophilus; Gribakin, 1979, Lethrus) is in favor of a universal membrane covering the microvilli. A fused rhabdom is difficult to analyze, since the PS of a given photoreceptor contributing to the rhabdom depends on morphological arrangement, crosssectional area, dichroic ratio of the microvilli, and the spectral sensitivities of all the rhabdomeres forming the rhabdom (Snyder, 1973), as well as on the twisting of the rhabdom (Grundler, 1974; Wehner et al., 1975; Wehner, 1976). Owing to optical coupling between the rhabdomeres of the fused rhabdom, the PS of a given photoreceptor may theoretically vary from a value much less than the dichroic ratio of a microvillus to a value greater than the dichroic ratio (Snyder, 1973). The most complete study of PS has been performed on the honeybee compound eye. Started in our laboratory as a morphological study more than 10 years ago (Gribakin, 1967a,b), it now shows that the low PS measured in honeybee photoreceptors (Menzel and Snyder, 1974) can be explained by twisting of the rhabdoms (see, for instance, Wehner, 1976), which rules out the hypothesis on electrical coupling suggested by Menzel and Snyder (1974). It appeared that only a small basal cell (the ninth cell), which seemed to be an ultraviolet light receptor (Gribakin, 1972), could display the high PS (up to 9) experimentally found by Menzel and Snyder (1974, also see Menzel, 1975a). Twisting of the rhabdom eliminates the filtering effect of distal ultraviolet receptors upon the ninth cell, and this makes the filter mechanism of polarized light perception in the bee suggested by Gribakin (1973) rather questionable, although it may operate in central rhabdomeres of dipterans (see also Snyder, 1973). If so, one must assume that in the ninth cell rhabdomere rhodopsin molecules are preferentially oriented to provide a dichroic ratio much higher than 2. Actually, a microvillus produced by rolling up the universal photoreceptor membrane must show a dichroic ratio of less than 2, that is, 1.64 (Gribakin and Govardovskii, 1975) or 1.67 (Laughlin et al., 1975). The theoretical limit for the dichroic ratio of a microvillus is 20 when ideal dipoles are all aligned along the microvillus, and about 6.5 in the case of nonideal dipoles (Laughlin et al., 1975). The most complete theory of the microvillar dichroism was advanced by Goldsmith and Wehner (1977). These authors also experimentally showed that the absorbing dipoles were aligned within k 50" of the microvillar axes in the crayfish Orconectes. In conclusion, we think that a systematic investigation of PS mechanisms in insects is needed not only to understand neural mechanisms of astroorientationin insects but mainly for a better understanding of cell membrane structure and function.

G. FINALREMARKS Light has made it possible for humans to unravel many mysteries of the structure of matter. Similarly, photoreceptor optics as applied to the cellular

159

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

mechanisms of photoreception in insects, provides a clue to the functional morphology of insect photoreceptors and, in particular, their photoreceptor membrane. However, both the functional morphology and structural lability of insect photoreceptors remains to be further investigated using methods of cell physiology. With this in mind we proceed to the electrical events resulting from the absorption of a photon by the photoreceptor membrane.

111. Electrical Basis for Insect Photoreception A. INTRODUCTION: THECOMFOUND EYEAS

A

VOLUME CONDUCTOR

Unlike a common neuron which transmits a signal as a local change in membrane conductivity traveling along the full length of its axon and manifested locally as a nerve impulse or spike, the arthropod photoreceptor transmits a signal electrotonically, that is, by a change in direct current flowing through its axon, and this has been unambiguously proved by Shaw (1972). In the transmission of a signal by direct current, one could consider the compound eye a volume conductor with many similar embedded electrical sources (i.e., cells, in both the biological and electrical sense) as well as with many similar electric (ionic) currents produced by these cells. Then the light effect on the compound eye extracellularly manifested as an electroretinogram (ERG) may be thought of as a disturbance of the electrical equilibrium peculiar to the eye in a state of dark adaptation. Thus we must first consider the potentials, currents, and resistances inherent in the eye in the dark and their changes during illumination. B. DC PARAMETERS OF THE COMPOUND EYEIN

THE

DARKAND IN

THE

LIGHT

1. Extracellular dc-Parameters of the Eye a. Dc Potentials. Our present knowledge of extracellular dc potentials (DCP) is much more limited than that of ERGS, and only several studies dealing with phenomenology rather than with the nature of DCPs have been performed (Burtt and Catton, 1964a,b; Cosens, 1967; Mote, 1970; Heisenberg, 1971). On reinvestigating DCPs in our laboratory, we have confirmed many previous findings and so are able to discuss the DCP problem with regard to our own experience. A typical DCP profile characteristic of the locust (Burtt and Catton, 1964a) is shown in Fig. 7. Two more curves are superimposed on the DCP profile. The first is the voltage difference A I/ measured between two electrode tips separated constantly in depth when penetrating an eye through which dc pulses have passed (Shaw, 1975); this curve reflects the axial resistance of the eye tissue. The

160

F. G. GRIBAKIN

V

4014aV

ERG mV

FIG. 7. Dc-characteristics of the locust compound eye. The solid line is the DCP profile V ( x ) , where x is the depth from the eye surface (Locusra migratoria, modified from Fig. 2a of Bum and Catton, 1964a). The dashed line is the voltage difference A V between two electrode tips separated by 115 p m in depth as 5-pA dc pulses pass through the eye from the cornea toward the basal membrane. The dashed-dotted line is the ERG peak amplitude. (Both curves modified from Shaw (1975) for Schistocerca gregaria). Top inset: Schematic sections of the Locusta (L) and Schistocerca (S) compound eyes according to Bum and Catton (1964a) and Shaw (1977). d, Dioptric apparatus (lens and cone); r, retinula; bm, basal membrane; lam, lamina; med, medulla; hc, hemolymph channel.

second curve is the ERG amplitude with depth (also, according to Shaw, 1975). In spite of the fact that the DCP profile has been measured in Locusta, and A V and the ERG in Schistocerca, there is not much difference in the size of the ommatidia in the two species, since, first, their basal membranes are located within a narrow range of depth (550-650 pm) and, second, the null point of the ERG is a clear indication of a distal border of the first negative voltage peak, and this is seen in Fig. 7. [For the null point of the ERG see, for instance, Fig. 2 in Cosens (1967); also, F. G. Gribakin and A. M. Petrosyan (personal observation) have found that the null point of the ERG is inevitably located at a level where the first peak reaches about half its amplitude, both in the locust and cricket]. According to Bum and Catton (1964a), the DCP profile of the eye may show three negative peaks coinciding with the first (lamina), second (medulla), and third (lobula) optic ganglia, and we have observed all three peaks in the locust, cricket, and cockroach (see Petrosyan, 1977a). The nature of all three peaks seems to be similar, and we deal only with the first. In brief, the properties of the first peak are as follows (Burtt and Catton, 1964a; Cosens, 1967; and personal

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

161

observations of Gribakin and Petrosyan). The first peak has no relation to the basal membrane, and its rising phase (i.e., the fall in potential in Fig. 7) coincides exactly with the distal border of the lamina. The highest peak value can be recorded only once-when the eye is first penetrated, though with a thick micropipet (a resistance of about 1 megaohm); during withdrawal, a second insertion, or when the electrode tip is left at the depth of the peak for several minutes, the peak displays strong damping, so that the potential profile can be totally smoothed out. However, if the eye is penetrated once again, but about 50 or 100 p m away, the negative peak is undisturbed. Thus the first peak is generated by local axial electrical sources situated at the outer border of the lamina; axial damage to the tissue from the electrode tip can abolish the peak only locally and the damaging effect does not spread radially to neighboring ommatidia. However, the peak can be greatly diminished over the whole eye by anoxia (to some extent, reversibly), while mechanical trauma to the lamina abolishes it irreversibly (Bum and Catton, 1964a). In general we recorded many normal three-peaked DCP profiles, though some of them showed no negativity near the cornea. Instead, a gradual increase in negativity along the full length of the ommatidium was observed. The effect of light on the DCP profile depends on where the reference electrode is placed. If it is in the insect body (in hemolymph), the DCP profile “seesaws” (Cosens, 1967) about the null point of the ERG profile, with more light-evoked negativity distal to the null point and with more positivity proximal to this point. In other words, the profile of the quasi-static ERG is the difference between the dark and light DCP profiles. With the reference electrode placed just beneath the cornea, the DCP profile can be shifted by light only positively, with no “seesaw” and with practically no change in potential until a depth of 400-500 p m is reached (in the locust). This means that using a corneal reference electrode no ERG can be recorded up to 400-500 pm. The effect of the reference electrode position on both the light DCP and ERG profiles could easily be explained by the presence of hemolymph channels just beneath the basal membrane, and these might serve as a ground point (the axis of the “seesaw”) when the reference electrode is located far away in hemolymph. The existence of these channels was postulated four years ago by A. D. Cherkasov of Moscow State University (personal communication) as an explanation of the ERG change in sign with depth, and experimentally the channels have recently been observed by Shaw (1977). The hemolymph channels appear to transfix the visual tract of the locust anterioposteriorly in between the basal membrane and the lamina where numerous trachea also have been found (Burtt and Catton, 1964a). Thus both the eye and the lamina have not only an excellent feeding system and oxygen supply, but also the ground point for ERG. There is no doubt that light-evoked changes in the DCP profile are produced by extracellular photocurrents of photoreceptors, as well as by the ‘‘response

162

F. G. GRIBAKIN

currents" of laminar neurons which respond to light with hyperpolarization (2ettler and Jhilehto, 1971), and the next question is what currents flow extracellularly within the compound eye in the dark and in the light. b. Dark Current and Photocurrent. In order to determine the direction of extracellular current flowing axially in the dark within the eye (i.e., parallel to its ommatidia), one must record the DCP profile V ( x ) ,where x is the depth from the eye surface. Then a profile of the axial electric field can be found as E ( x ) = -dV(x)/dx, where the minus sign means that the current is defined as a flux of positive charges forced to move by the electric field E ( x ) which flows in the direction in which V ( x ) decreases. The calculated profile E ( x ) (Fig. 8) clearly shows that the extracellular axial current directed by E (x) has to reverse its sign somewhere in the depth of the eye. (Note that E ( x ) does not depend on a shift of the whole V ( x ) curve along the V axis, hence does not depend on where the reference electrode was placed when V ( x )was recorded.) It appears that, even in the dark, three current loops exist within the eye zone considered (Fig. 8). If distal negativity is absent, the first loop disappears and the second one spreads, covering the whole retinula and possibly the cone cells. Since every current loop is to be closed, the photoreceptors and neurons of the lamina serve as the return - 9 1

, & ' I'

"

.+. -i. -l -- -- .---,[4 - -., ..--*.,..U' E

5-

"

"

1 -*-'

I-,

V

mV

FIG.8 . Dc characteristics of the locust compound eye. The solid curve is the smoothed DCP profile V ( x ) , as in Fig. 7. The dashed curve is the electric field E ( x ) given as a derivative of V ( x ) with respect to depth from surface x taken with a reverse sign since E(x) = - dV(x)/&. The graph of E ( x ) characterizes the direction of the current flowing along the ommatidia but outside them. Top inset: Three possible current loops in the dark resulting from the sign of the graph E ( x ) obtained.

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION I

'

'

' 1

f

'

'

'

'I

-- -- - --

I

- - - I

'

' I '

I

'

'

163

I

It *---\--+.:-,\--*.-I

c-

I _ _ _ _ -

- 1 5 ERG

"wires" for the extracellular currents and probably as their sources. A quantitative evaluation of the dark currents in the compound eye demands an accurate knowledge of tissue resistance, however, their approximate values may be given as follows. Near the basal membrane the electric field E is about 0.13 V/cm (taken from Fig. 7 or 8). Taking into account that pigment cells may serve as a conductive medium (which is thought to be true for the glial processes in the brain; see Ranck, 1963) with a specific resistance p of about 80-100 ohm-cm, we obtain the dark cumnt density as j = E / p = 1.3 x Alcm2. At the basal membrane the axonal bundle of a locust retinula has a diameter d of about 4 pm, and the mean distance D between the axes of adjacent retinulas is about 8 p m (F'etrosyan, 1977a). Then a conductive extraretinular cylinder has the crosssectional area S = d 4 (D2- &) = 3.8 X lo-' cm2, and the extraretinular dark current i d per retinula about the basal membrane is i d = j X S = 5 X A, that is, 0.5 nA per retinula (or, divided by eight cells, 0.06 nA per photoreceptor). A whole eye consisting of 8000 ommatidia (locust) would show a dark current of 4.0 pA at the basal membrane. In the light, the extracellular dark current changes, and this change can be termed the extracellular photocurrent. The additional voltage drop produced by photocurrent flowing through extracellular (more exactly, extraretinular) resistance can be measured as an ERG. From the ERG amplitude profile with depth (Fig. 7), one can determine that extracellular photocurrent underlying the ERG flows toward the cornea, that is, opposite the dark current. In other words, light diminishes the dark current (Fig. 9). If the specific resistance of the lamina were of the same order of magnitude as that between the cornea and basal membrane, one could expect maximal photocurrent density somewhere in the lamina where

164

F. G . GRIBAKIN

FIG10. A possible current path for pulses passed througb the eye to measure its resistance. The current electrode A is at the cornea, and the current electrode B is in the hemolymph. The resistance profile can be obtained in two ways: (a) two electrodes with tips separated longitudinally move from the cornea along the x-x line in order to measure the potential difference A V arising in tissue when current pulses pass from A to B (see Shaw, 1975); (b) the reference electrode is located near the electrode A, and the probe electrode is inserted by steps along the x-x line (Cribakin and Petrosyan, hitherto unpublished). Both potential profiles reflecting the eye resistance are shown qualitatively as curves a and b. R, Retinula; PC, pigment cells; L, lamina; H, hemolymph; bm, basal membrane.

the electric field of the ERG is maximum (Fig. 9). A maximum ERG amplitude, and consequently zero axial photocurrent, may indicate that these are the terminals of axons through which all the photocurrent produced by photoreceptors flows outward in the synaptic region. A quantitative evaluation of the ERG photocurrent cannot be made, since the number of stimulated ommatidia is not known as a rule from the studies available. However, from the fact that dark current and photocurrent produce similar voltage drops in the extracellular medium of the eye in routine experiments with ERG recordings, one can infer that the photocurrent at the basal membrane is within the same limits as the dark current, that is, about 0.1 nA per receptor (though it strongly depends on the light intensity). Comparisons between the dark current loops (Fig. 8) and the photocurrent loops (Fig. 9) show that it is the second dark current loop which shares a current path with the photocurrent at the retinular level, and this indicates that the receptor signal can be interpreted as a modulation of the dark current within the second loop. Thus the principal problem to be solved concerns what emf sources are responsible for this second loop current. Of course, not only photoreceptors but, equally, neurons of the lamina may appear to be involved. c. The Resistance Profile. The actual resistance of the compound eye is not known exactly, and only two attempts to determine it have been made. Shaw (1975) measured the voltage drop A V ( x ) between two electrodes with tips 115

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

165

p m apart in depth when 5-pA current pulses were passed axially through the eye of a locust (Fig. 7). The resistance profile found by Shaw can easily be interpreted if again the existence of hemolymph channels (Shaw, 1977) is taken into account. Assume that the paired electrodes are inserted into the eye along the x-x line as shown in the model in Fig. 10. If the basal membrane is a resistance barrier (Shaw, 1975), the current will flow from the pigment cells and extracellular space into the photoreceptors, and this will cause a decrease in AV with depth. However, as soon as one of the electrodes penetrates a hemolymph channel, a sharp rise in the potential AV will be measured. Then, when it is inserted deeper, A V must change its sign, since the current pulse in the lamina has the opposite direction between the outer border of the lamina and the terminals of retinular axons. When the electrode is inserted still deeper, A V becomes positive again (cf. Figs. 10 and 7). However, if the reference electrode is placed near the current electrode A (Fig. lo), and the probe electrode is inserted by steps into the eye, a different profile like that shown in Fig. 10 (curve b) can be recorded (F. G. Gribakin and A. M. Petrosyan have used this second method in our laboratory; see Fig. 11). Then, a sudden jump in potential at the basal membrane reflects the penetration of a hemolymph channel but, when the electrode reaches the lamina, the potential difference thus measured drops again. The value of the postbarrier potential divided by the pulse current might be taken as the approximate resistance of all the photoreceptors connected in parallel. Indeed, if the axons are the only conductors inside the eye connected to the inside of the lamina (for current

FIG.1 1 . The voltage drop V between a corneal reference electrode and a probe electrode inserted stepwise into the eye when current pulses have passed through the eye as shown in Fig. 10. Note that the voltage drop appeared to be nearly the same in three preparations beyond the resistance barrier. This voltage divided by a 5-c~.A cumnt pulse gives a cornea-lamina resistanceof about 4-5 kilohms. (F. G . Gribakin and A. M. Petrosyan, hitheno unpublished; Locusta preparations 17-L, 18-L and 19-L.)

166

F. G. GRIBAKIN

pulses which then flow out of the lamina through its envelope), then all the photoreceptors have plasma membranes (or input resistances) connected in parallel. Taking the number of photoreceptors in the locust eye as about 60,000 and the input resistance of a single photoreceptor as about 10 megaohms (Shaw, 1969a), we obtain the eye resistance for a uniform current pulse of 170 ohms. Then, taking the average axon length equal to 200 p m and its diameter 1 pm, we obtain a single axon resistance, 100 megaohms (with a specific axoplasm resistance of 100 ohm-cm). Further, if connected in parallel, all 60,000 axons would show a resistance of 2 kilohms. Subtracting this value from the total cornealamina resistance (Fig. l]), we obtain 2-3 kilohms which are probably due to synaptic membranes. As a rule, a potential difference of 10-15 mV exists in the dark between the extracellular interior of the eye and that of the lamina. This voltage difference is probably due to total dark current flowing through the axons, and again we obtain the total dark current of the eye equal to 2.5 p A , taking a parallel axonal resistance of 4 kilohms and a voltage difference of 10 mV (cf. 1.5-2 p A in previous paragraph). d. Conclusion. The compound eye must be considered a volume conductor showing a complicated pattern of extracellular DCPs and currents existing even in the dark. Nothing is known at present about the cellular mechanisms responsible for this dark electrical activity which does not seem to reflect exclusively the metabolism of the compound eye but undoubtedly underlies its principal physiological function. 2. Intracellular Dc Parameters of the Photoreceptors and Accessory Cells a. Resting Potentials. Resting potentials of photoreceptors have not been measured systematically, though a precise knowledge of the potential difference between the inside of a photoreceptor and its extracellular environment is necessary for an understanding of the ionic mechanisms of both dark emf’s and the photoexcitation process. To fill the gap, in part, we have made measurements of resting potentials of the photoreceptors and the cone cells in three insect species and have found that the resting potentials, first, are quite different in the photoreceptors and cone cells of the same species (though both cells are surrounded by the same extracellular medium) and, second, differ within the same cell type in different species (Gribakin and Petrosyan, 1979; see Table IV). Relatively low resting potentials of photoreceptors might be due to a slight permeability of their membranes to sodium in the dark, as Fulpius and Baumann (1969) have suggested in order to explain the deviation of the drone bee photoreceptor resting potential from the Nernst equation produced by a change in external potassium. Also, appreciable sodium permeability might explain why resting potentials of photoreceptors is lower in species with a higher sodium level in the hemolymph (Gribakin and Petrosyan, 1979). Here two possibilities are considered to explain

167

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION TABLE IV RESTINGPOTENTIALS OF ~OTORECEFTORS AND CONECELLS~ ~

~

~~~~~~~~

Resting potential (mV) Photoreceptors Insect species Locusra rnigratoria Periplaneta americana Gryllus domesticus

Cone cells

Extreme values

Mean T SD

Extreme values

Mean fSD

15-75

42?13(n=40)

20-95

6 0 k 16(n = 51)

10-65

32?lO(n=72)

30-70

55 k 10(n = 56)

5-45

23 f 1 1 (n = 45)

15-85

49 4 16(n = 36)

the relatively low photoreceptor resting potential, and the photoreceptors of the house cricket G. domesticus are taken as an example. The first possibility is based on two statements: (1) the lower resting potential of the photoreceptor is due totally to the noticeable permeability of its plasma membrane to sodium ions, and (2) extracellular concentrations of sodium and potassium are the same as those in the hemolymph. An alternative explanation rests upon the assumption that sodium permeability in the dark is negligible and that statement 2 is not true since the extracellular ionic content in the nerves and ganglia of many insects greatly differs from that of their hemolymphs (see review of Treherne and Pichon, 1972). For the first possibility, the sodium/potassium permeability ratio may be calculated from the Goldman equation:

Em = 58 log,,

W+I,+ b"a+I,

[K+],

+ b[Na+Ii

where E m is the average resting potential measured (see Table IV); [K+],, [K+II, "a+],, and [Na+II are the concentrations (or, to be more exact, activities) of potassium and sodium inside and outside the photoreceptor; and b is the sodiudpotassium permeability ratio. The potassium concentration in the cytoplasm [K+Ii is taken equal to 120 m M as revealed by x-ray microprobe analysis (Gribakin et al., 1976; Burovina et al., 1978). The sodium content [Na+Ii is taken equal to 55 mM, since flame photometry data have given a concentration of 55 mM as the mean for the whole eye (Petrosyan et al., 1977), while electron cytochemistry has demohstrated sodium pyroantimonate precipitate, mainly in the photoreceptors (Petrosyan, 1977a,b). "a+], and [K+], are taken as equal to

168

F. G.GRIBAKIN

208 and 8 mM for cricket hemolymph (Petrosyan et al., 1977). For the mean E = 23 mV (Table IV), the value of b = 0.21 is obtained which appears to be too high in comparison with the smaller b value postulated from the experiments of Fulpius and Baumann (1969) in which the total removal of external sodium resulted in only slight hyperpolarization of the drone bee photoreceptor (about 5 mV). For the second explanation, “a+], and “a+]* can be totally ignored, since sodium permeability in the dark is assumed to be negligible. Then, a similar potassium content in the extracellular medium might be expected when calculated from the resting potentials of either cone cells or photoreceptors as both having the same extracellular surroundings. From our x-ray microprobe data, averaged [K+] is 170 mM in the cone cells and 120 mM in the cytoplasm of a photoreceptor (Burovina et al., 1978). Using &. (12), with b = 0, one can find that [K+], for the cricket cone cell (resting potential 49 mV; see Table IV) and the photoreceptor (resting potential 23 mV) are 24 and 48 mM, respectively. This 2-fold difference in calculated values of [K+], is probably due to many reasons and, particularly, to the fact, that we used concentrations, not activities, here which are still unknown. There is much evidence that neither photoreceptors of insects (drone bee, Fulpius and Baumann, 1969) nor their neurons (Treherne and Pichon, 1972) are potassium electrodes, since their E m versus [K+], plots show a lesser slope than that calculated using Nernst’s equation. However, H. M. Brown (1976), using potassium-sensitive intracellular microelectrodes, showed that, in Balanus photoreceptor, [K+] i increased with growth in external potassium concentration [K+],, and this accurately explained the difference between the experimental and theoretical Em versus [K+], plots. The principal conclusion of H. M. Brown was that the plasma membrane of the balanus photoreceptor does behave as a potassium-sensitive electrode. In conclusion, we think that the relatively low resting potentials of insect photoreceptorsmay serve as an indication of a higher extracellular potassium content in the eye rather than of their noticeable permeability to sodium in the dark. It should be noted that the higher resting potentials of the cone cells make them similar to glial cells which inevitably display resting potentials higher than those of adjacent neurons; this is thought to be due to their possible role as a mechanism controlling the extracellular potassium level in nerve tissue (see review of Somjen, 1975). b. Dark Current of a Single Photoreceptor. In Section III,B, 1 ,b extracellular axial dark current in the compound eye has been described and the photoreceptors were considered the return wires. Practically, the reverse is more convenient with a photoreceptor as the direct conductor through which two intracellular dark currents flow; the first flows proximally in its distal region, and the second distally in its proximal region, including the whole axon (Fig. 8). To avoid misunderstandings connected with the terms “proximal” and “distal,

”

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

169

which are related to both region and direction, we term these currents the receptor dark current (with its receptor loop) and the axonal dark current (with its axonal loop). Nothing can be said at present about the cellular origin and ionic carriers of these currents, though the receptor current might be related to an ionic pump mechanism which seems to be inherent in the distal third of the photoreceptor where the bulk of its mitochondria is located (seee.g., review of Gribakin, 1969~).The axonal dark current can be considered to be generated by the electrical source responsible for the first DCP peak, and consequently located in the lamina. Thus the axonal dark current does not seem to be due to photoreceptor electrical activity but probably to that of the first neuron in the lamina. From this an interesting speculation arises that the receptor dark current reflects mainly metabolic functions of the photoreceptor necessary to provide its high absolute sensitivity (and, probably, the ionic gradient needed), while it is the first neuron which continuously sends its “inquiry current” through the receptor axon to determine the state of the receptor. If so, the receptor might modulate this “inquiry current” upon absorption of a photon, and this modulation might be what we know as the receptor signal. Only further studies will c o n f i either the above speculation or the more prosaic conclusion that the axonal dark current is no more than a simple background phenomenon having nothing in common with the receptor signal. At any rate, at present we can confirm the presence of two intracellular dark currents in a photoreceptor and calculate the axonal dark current to be 0.06 nA in the locust (Section III,B), which corresponds to about 4 x 108 unit charges per second. A voltage drop produced by the dark current through the axon can be as small as 6 mV, taking the dark current as 6 x lo-” A and the resistance of the axon as 10’ohms. A voltage drop characteristic for the synaptic membrane of the locust photoreceptor in the dark is obtained as the total dark current of the eye (4 pA) times the total synaptic membrane resistance (2-3 kilohms), which gives 8-12 mV, negative inside.

C. PHOTORESFONSES AND THEIR CELLULAR MECHANISMS 1. Photoresponses

Several types of photoresponses inherent in the compound eye can be recorded intra- and extracellularly. First, intracellular miniature potentials, discrete waves, or quantum “bumps,” according to different terminology. We use the latter term “bumps,” which is more felicitous (cf. Levinson, 1972) and concise, though “miniature potentials” apparently is more correct (e.g., Tsukahara and Homdge, 1977). Second, also intracellular, is the receptor potential, and this term is thought to be more suitable in comparison with the term “generator potential” used, for instance, by Fuortes and O’Bryan (1972a,b); in our opinion,

170

F. G . GRIBAKIN

the term “generator potential” reflects rather an active, regenerative process, hence it should be used to designate the receptor repetitive spikes, if any. In turn, four components of the receptor potential are, consecutively, the spike, transient, plateau, and prolonged depolarization afterpotential (PDA) (Goldsmith and Bernard, 1974; Wulff and Mueller, 1975; Hamdorf and Razmjoo, 1977). Third, the extracellularly recorded photoresponse is the ERG which occurs as a voltage drop produced by net photocurrents of the receptors in the extracellular medium of the eye. In this section we consider these photoresponses only from the viewpoint of their cellular origin.

2. Quantum Bumps Quantum bumps in arthropod photoreceptors are thought of as electrical manifestations of single photon events (see, for review, Fuortes and O’Bryan, 1972b), but whether or not these are one-to-one transformations is still not clear. Rare bumps are clearly seen even in complete darkness (Yeandle and Spiegler, 1973, for Limulus; Tsukahara and Horridge, 1977, for the locust), and with an increase in illumination they fuse to give rise to the receptor potential. If considered voltage noise during the receptor potential, bumps correspond to a standard deviation of 1.55 mV in the cockroach, 0.7 mV in the fly, and 0.6 mV in the bee (Smola, 1976). On the assumption that one light quantum triggers a mechanism generating one bump, Levinson (1972) has defined the energy gain as bump energy related to photon energy and has obtained the value of lo6 for Limulus photoreceptors. According to another definition, the gain is expressed as an increase in “particles,” or current carriers, and in this sense the gain in Limulus photoreceptors appears to be about 6 x lo8 unit charges per absorbed photon (Levinson, 1972). For the locust, the energy gain can be approximately evaluated, taking the bump amplitude as 3 mV, the duration of the bump as 0.1 second Psukahara and Horridge, 1977), and the cell input resistance of 10 megaohms (Shaw 1969a) as 2.5 x lo5 per photon with a wavelength of 500 nm; this value is of the same order of magnitude as that in Limulus. The amplification thus found favors the presence of a yet unknown process which has to intervene between photon absorption by the photoreceptor membrane and a change in potential across this or, perhaps, the plasma membrane of the cell. The transduction process is seemingly mediated by one or more intracellular transmitters. Pak et al. (1976) demonstrated that, in norpd Drosophila mutants with phototransduction blocked totally or in part, neither rhodopsinmetarhodopsin photoisomerizationnor bump-generating machinery had been disturbed. They concluded that the norp-A mutation affected a step which preceded bump production. This step may, for example, be involved in the release of a substance which in turn produces quantum bumps (pak et al., 1976). The fact that bumps remain rather a long time after a strong light is switched off (and consequently when there are no light quanta to be absorbed) may also be in favor

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

171

bump generation through an intracellular transmitter produced by light in excess, which continues to act for some time in the dark (Minke et al., 1975; Tsukahara and Horridge, 1977). According to Martinez and Srebro (1976), a decrease in extracellular calcium leads to an increase in the latency of the bumps and their variability (this effect is probably due to a subsequent decrease in intracellular calcium). Lowering the temperature has given the same result. However, since the quantum efficiency of the process remains the same in both cases, Martinez and Srebro have concluded that calcium probably affects the rate constant of one of the intermediate reactions. From the fact that, the higher the extracellular calcium, the lower the amplitude of the bumps (Millecchia and Mauro, 1969; Martinez and Srebro, 1976), one can assume that another function of calcium is to reduce bump amplitude in the light, for bumps are known to be smaller in the light (Dodge et al., 1968; Fuortes and O’Bryan, 1972b; Srebro and Behbehani, 1974, for Limulus; Tsukahara and Homdge, 1977, for the locust). On the contrary, in trp mutants of Drosophila bumps show no decrease in amplitude in the light, and so Minke et al., (1975) suggested a lower efficiency of bump production in the light. Thus there is no clear distinction so far between a diminution of the electrical response per photon absorbed at higher intensities (i.e., a fall in bump amplitude) and a decrease in the relative number of photons (divided by incident) per rhodopsin molecule provided by cellular mechanisms of absolute sensitivity control (see Section II,D, and Tsukahara and Horridge, 1977). The last stage of bump generation is generally beIieved to be the activation of sodium channels by a hypothetical transmitter, but whether the number of the transmitter molecules in one transmitter quantum is a constant and, if so, the number of channels it can activate simultaneously are not known (Tsukahara and Horridge, 1977). It also remains to be clarified whether bumps really fuse to form the receptor potential or the two electrical responses are based on different cellular mechanisms, even with different cellular membranes involved, so that bumps are simply masked by the larger receptor potential at higher intensities. At any rate, the trp mutant of Drosophila has shown a “bumpy” receptor potential with no fusion of bumps (h4inke et al., 1975). 3 . Receptor Potential a. Spike. An initial spike has been found to contribute to the receptor potential in many arthropods (see review of Goldsmith and Bernard, 1974; Wulff and Mueller, 1975; Baumann, 1975), though it does not seem to be a requirement in insects. For example, we have observed the spike in the beetle k t h r u s but never in the locust (neither have Tunstall and Honidge, 1967) or the cricket. In the drone bee the spike is due totally to a regenerative inflow of sodium ions into the photoreceptor, since it is abolished in a sodium-free solution (Fulpius and Baumann, 1969) and can be blocked reversibly by tetrodotoxin (Baumann, 1968,

172

F. G . GRIBAKIN

1975). Upon withdrawal of the recording microelectrode from the cell, the spike does not change its polarity, unlike the other components of the receptor potential, and this means that the spike is generated by cellular elements other than the plasma membrane (Baumann, 1968, 1975). According to Naka and Eguchi (1962), the spike originates in the axon, though the photoreceptor membrane might be a better candidate for this response (Wulff and Mueller, 1975, for Limulus). If so, the fact that the spike is the earliest of the light-evoked ionic processes in the photoreceptor might result from a sodium permeability increase in the photoreceptor membrane itself, whereas both the transient and plateau develop as the result of an intracellular transmitter action upon the peripheral plasma membrane of the photoreceptor. b. Transient, Plateau, and Related Cellular Mechanisms. The plateau is the third component of the receptor potential on the time scale (after the spike and transient), however, it is the first resulting from bump fusion at intensities which still remain low in order to initiate spike and transient generation. Only at comparatively high intensities is the plateau preceded by a transient and, when a firing level of depolarization is reached, the transient in turn may be preceded by a spike. Wulff and Mueller (1975) hold that in the lateral eye of Limulus the plateau originates in the photoreceptor membrane of the rhabdomere, as the spike does, while the transient is generated in the plasma membrane of the receptor. In favor of this is the fact that the reversal potential for the spike and plateau is the same-about +16 mV, while that for the transient is only +4 mV. From the positive sign of the reversal potential, Wulff and Mueller (1975) have deduced that not only the spike but both the transient and the plateau are also due mainly to sodium influx in the light. Apart from the Limulus lateral eye, a sodium permeability mechanism seems to operate in Limulus ventral photoreceptors, the barnacle ocellus, the compound eye of the crayfish and hermit crab, and the compound eye of the drone bee (for review, see Wulff and Mueller, 1975). A controversy, however, has developed concerning the well-known fact that photoreceptors of arthropods are capable of maintaining the receptor potential, though it is markedly decreased, in sodium-free solutions (Fulpius and Baumann, 1969; Baumann, 1975; Wulff and Mueller, 1975); and in general this ability of arthropod photoreceptors is not unique among the sensory end organs of animals (see, for instance, Brown and Ottoson, 1976). In order to resolve this contradiction, an assumption has been put forward that even in a sodium-free solution a significant amount of sodium is preserved close to the plasma membrane of the receptor (Stieve, 1964, for arthropod photoreceptors; Ottoson, 1964, for muscle spindles). This assumption has proved to be very useful, and sodium pump activity has been suggested as a factor responsible for maintaining the sodium concentration gradient across the photoreceptor cell plasma membrane even in a sodium-free solution (Wulff et al., 1975, for Limulus). However, Wulff et al. (1975) note that the inherent weakness of their hypothesis is that “rigorous proof

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

173

rests with the demonstration of sodium concentrations in the various compartments associated with the Limulus lateral eye retinular cells, the techniques for which are not at hand. ” Indeed, the principal demand of the above assumption is that sodium be stored in either accessory cells of the eye (pigment cells or cone cells) or in the photoreceptors themselves. According to our measurements, the sodium content of the insect compound eye is relatively large, about 55 meqkg wet weight in the house cricket and 14 meqkg wet weight in the locust (Petrosyan et al., 1977). When tested by electron cytochemistry using a potassium pyroantimonate technique to precipitate sodium, the bulk of the deposit was found in the photoreceptors and not in the other cells of the eye (Petrosyan, 1977a,b, for cricket)’. X-ray microprobe analysis has shown the sodium content in cricket photoreceptors to be 40-70 mM (Gribakin et al., 1977; Burovina et al., 1978). The intracellular sodium activity has not been measured in insect photoreceptors, but in those of the barnacle it appears to be about 10 mM in the dark and 18 mM after illumination with bright light (Brown and Comwall, 1975b; Brown and Ottoson, 1976) or, according to other data, about 25 mM in the dark with no more than a 5-mmole increase in the light (Brown, 1976). Thus we suggest that the photoreceptors themselves are capable of storing sodium, hence they can supply their own ionic pump with internal sodium for a prolonged time even in a sodium-free solution (Gribakin et al., 1977; Petrosyan, 1977a). The “sodium mechanism, consequently, can be accepted for insect photoreceptors, and this may underlie generation of both the transient and the plateau. It should be noted that the concepts of transients and plateaus are convenient but formal descriptions of the receptor potential components. Indeed, the drop from the transient to the plateau is mediated by intracellular calcium. After intracellular injection of EGTA the transient fails to drop to the plateau, and the receptor potential remains at the transient level as long as the light is on (Lisman and Brown, 1975; Bader et al., 1976). These experiments, as well as those with intracellular injection of sodium and calcium (Lisman and Brown, 1972; Bader et al., 1976), allow the transient-plateau transition to be interpreted as a result of the action of a calcium-mediated feedback intended to provide absolute sensitiv”

‘According to Petrosyan (1977a,b), all deposits should be related to sodium (not calcium), since (1) all sodium in the cricket photoreceptors is accesible to water, while calcium is practically not (see Burovina et al., 1978), and (2) sodium concentration as well as its activity in the cytoplasm, (10-50mM, see text for further details) is several orders of magnitude higher than that of calcium (no more than M ,see Borle, 1973; Bygrave, 1978). Thus calcium could hardly be responsible for the abundant deposit found in the cricket photoreceptors. Similarly, demonstration of calcium localization in the drone bee photoreceptors by Perrelet and Bader (1978) using pyroantimonate technique seems to be doubtful for these reasons (in addition, in the latter experiments, the treatment of the eye with the pyroantimonate solution had been preceded by prolonged fixation with glutaraldehyde solution; this procedure inevitably led to a redistribution of elements under study and possibly to their fractional or even total removal from the tissue; x-ray microprobe analysis in this case might indicate calcium still bound in vivo, not necessarily deposited by the reagent).

174

F. G . GRIBAKIN

ity control at the level of the receptor potential. In this interpretation, oscillations of the receptor potential often observed during the transient-plateau drop at high-intensity stimuli (e.g., see Fig. 12 in Fulpius and Baumann, 1969) are identical to those arising in electrical and mechanical oscillatory systems with nearly critical damping. Thus the plateau level of the receptor potential is a result of the interaction of two opposing ionic mechanisms: first, the light-induced sodium influx which depolarizes the cell membrane to the transient level and, second, a calciummediated mechanism (probably triggered by the former process) which tends to reduce this depolarization (antitransient repolarization). This interaction using a calcium-mediated feedback provides a wider dynamic range of receptor functioning at the plateau level in comparison with the transient level (see Fig. 2 in Fulpius and Baumann, 1969; Fig. 2 in Stieve et al., 1976). The light-induced calcium increase in the photoreceptor directly demonstrated by Brown and Blinks (1974) in Limulus ventral photoreceptors using aequorin may be due to the liberation of calcium ions from intracellular stocks activated by a small light-induced increase in intracellular sodium (Bader et al., 1976). In the arthropod photoreceptor calcium may be stored intracellularly, not only in mitochondria (see, e.g., Borle, 1973) but also within the pigment granules where it does not seem to be directly accessible to water (Burovina et al., 1978). Consequently, both mitochondria (Carofoli et al., 1974; Bader et al., 1976) and pigment granules might serve as intracellular calcium reservoirs (or depots) [Burovina et al., 1978; cf. the light-induced calcium release from the pigment granules of Aplysia neurons reported by Brown et al. (1975)]. In this respect, the migration of mitochondria and pigment granules to the rhabdom in the light considered previously to be the optical mechanism of absolute sensitivity control (Section II,D) may have a double function, the second purpose being to provide a photoreceptor zone of the cell with additional calcium to prevent a fatal increase in permeability to sodium under bright illumination. The PEC can hardly be considered an ionic depot, as suggested by Bader et al. (1976), because of its watery content (Section II,D,S,c). Moreover, we failed to find in the PEC an increased amount of any cation; on the contrary, potassium has a concentration in the PEC nearly half that in the photoreceptor cytoplasm (Gribakin et al., 1976; Burovina et al., 1978). However, participation of the PEC in the transduction process cannot be excluded, taking into account the potassium gradient on its membrane. In conclusion, the transient and plateau-principal components of the receptor potential-need to be interpreted as the algebraic sum of continuous depolarization (with the amplitude of the transient) produced by sodium influx and a subsequent opposing process-more slowly developing calcium-mediated repolarization. Both the transient and plateau probably originate in the plasma

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

175

membrane of the photoreceptor, but not in its rhabdomere, and are triggered by a still unknown intracellular transmitter. c. Prolonged Depolarization Afterpotential. After the strong or prolonged light stimulation (especially with monochromatic light) ceases the plateau fails to fall sharply to a resting potential. Instead, the depolarizing afterpotential is seen to last from several seconds to many hours depending on the light stimulus (wavelength, intensity, time) and the species (Baumann and Hadjilazaro, 1972; Nolte and Brown, 1972; Hochstein et aE., 1973; Brown and Cornwall, 1975a,b; Hamdorf and Razmjoo, 1977; Tsukahara et al., 1977; Hillman et al., 1977). This effect has been termed prolonged depolarization afterpotential (PDA). A mechanism generating a PDA was first suggested by Baumann and Hadjilazaro (1972); “The retinula cell produced some unknown substance which, by acting on its membrane, leads to depolarization. Strong or prolonged illumination might lead to the accumulation of this substance and, owing to its relatively slow disappearance at the end of the flash, to a prolongation of the response into the dark.” Using the early receptor potential (ERP) as a measure of the rhodopsin concentration, Minke et al. (1973) and Hochstein et al. (1973) demonstrated the existence of two thermally stable states of rhodopsin in the barnacle eye, as well as a correlation between metarhodopsin formation and the presence of PDA, so that PDA is restrained by the amount of metarhodopsin produced by a light flash. Since PDA can easily be abolished by a flash with a wavelength corresponding to the metarhodopsin absorption maximum, to the early suggestion of Baumann and Hadjilazaro (1972) can be added the idea that removal of an unknown substance accumulated during rhodopsin conversion is prolonged when the pigment is in a metarhodopsin state Psukahara et al., 1977). An ionic mechanism underlying PDA is thought to be the same as that underlying the plateau, that is, a prolonged increase in sodium permeability, since both responses display the same dependence on ionic environment and temperature (Baumann and Hadjilazaro, 1972, drone bee; Brown and Cornwall, 1975b, barnacle). The role of calcium in PDA fading or abolition is still unknown (Brown and Cornwall, 1975b). Like the ERP, PDA does not seem to arise in natural light environments (see Hochstein et al., 1973) and, for this reason, has no real physiological function or significance; but it can be considered a tool for the study of visual pigments and their conversions triggering the ionic mechanisms that generate the receptor signal. 4. Photoresponses of Accessory Cells of the Eye a. Pigment Cells. Pigment cells are capable of responding to light with a depolarization which develops slower than the depolarizing response of the photoreceptors (Bertrand et al., 1972; Baumann, 1975). The resting potential of the pigment cells in the drone bee is lower than that of the photoreceptors (45 mV

176

F. G. GRIBAKIN

average to 50 mV average in the photoreceptors) and depends mainly on the potassium concentration in the extracellular medium. The response to light decreases when the potassium concentration in the extracellular medium is increased. The resistance of the pigment cell plasma membrane (or input resistance) does not change in the light, unlike that of a photoreceptor. Because of this, the pigment cell response is thought to be due to an increase in extracellular potassium leaving the photoreceptors in the light (Bertrand et al., 1972; Baumann, 1975). Pigment cells can be thought of as regulators of the extracellular ionic concentration (Baumann, 1975), and this function may appear similar to that suggested for neuroglia (see Somjen, 1975). b. Cone Cells. Like pigment cells, cone cells respond to illumination by slow depolarization lasting for 15-20 seconds after the light goes off (Vishnevskaya and Mazokhin-Porshnyakov, 1969; Cherkasov e f al., 1976; Vishnevskaya e f al., 1977), and this response is also assumed to be due to an increase in extracellular potassium in the light, as in the case of pigment cells (Vishnevskaya et al., 1977; Petrosyan, 1977a). An ability of cone cells to respond to light has also been demonstrated by electron cytochemistry. Petrosyan (1977a,b) found that numerous granules of sodium pyroantimonate inherent in cricket cone cells in a state of dark adaptation disappeared from these cells when they became light-adapted. However, both the mechanism and significance of this response are still obscure. Spectral efficiency characteristics of cone cells coincide with those of the whole eye, as measured by ERG, and can change during color adaptation in the grasshopper Teftigonia which has at least two spectral types of photoreceptors (Vishnevskaya and Mazokhin-Porshnyakov, 1972; Cherkasov et al., 1976). This finding favors the above assumption that the cone cell response is triggered by the activity of the photoreceptors. Similar to what has been assumed for the ion-controlling function of pigment cells, cone cells might also be involved in this regulation, and four cone processes going along the photoreceptors and terminating at the basal membrane as pigment-filled expansions (Gribakin, 1967, 1975; Wachmann e f d.,1973; Polyanovskii, 1976) may form a morphological basis for this function.

5 . Electroretinogram The ERG is defined as a light-evoked potential difference recorded between two electrodes one of which is placed just beneath the cornea while another is inserted elsewhere in the insect body. The ERG was the first electrical response of the compound eye to be recorded, and the history of its investigation as well as more details on ERG interpretation can easily be found in the literature (Mazokhin-Porshnyakov, 1969; Heisenberg, 1971; Goldsmith and Bernard, 1974). Being a mass response the ERG is produced by all the extracellular currents flowing in the illuminated eye and generated mainly by photoreceptors and laminar neurons. Consequently, the two principal components of the ERG

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

177

are the receptor and lamina responses. The ease with which the ERG can be recorded is rather a warning to workers of the difficulties arising in ERG interpretation in terms of lightevoked changes in extracellular photoreceptor dark currents or, to be more exact, in terms of cellular mechanisms of insect photoreception. At any rate, many questions concerning ERGS clearly formulated by Heisenberg (1971) are still being investigated. D. THEROLEOF COMPARTMENTALIZATION The role of compartmentalizationwas first noticed by Heisenberg (1971), who suggested that the compound eye be considered an insect body compartment. This is in line with the ideas developed by Treheme and Pichon (1972) that the ionic composition of the extracellular medium of an insect nerve or ganglion should be stabilized to provide reliable functioning of the nerve system independently of the inconstant life conditions of the insect. Similarly, the extracellular medium of the eye can in no way be considered identical to the hemolymph and needs to be specially studied. The discovery of the resistance barrier (Shaw, 1975) and diffusion barrier (Shaw, 1977) in the compound eye, as well as direct studies of ionic composition of the eye and its elements (Gribakin et af., 1976, 1977; Petrosyan 1977a,b; Petrosyan et al., 1977; Burovina et al., 1978), have clearly shown that the compound eye is really a compartment of the insect nerve system which in turn contains “second-order” compartments (i.e., different cells) among which photoreceptors are strictly compartmentalized so that even their potassium is far from being uniformly distributed in the cell (Gribakin et al., 1976; Burovina et al., 1978). A correlation between ionic and morphological compartmentalizationmay favor various cellular sources of different receptor potential components and perhaps provide the clue to understanding the cellular mechanisms of receptor signal generation.

IV. Conclusion In this section several important principles and ideas following from this account are briefly summarized. Adaptation of the insect eye to the dark is an active process showing a continuous demand for energy to create and maintain a highly efficient optical structure ensuring perfect absorption properties. In a poor light environment the insect photoreceptor is able to operate as a photon counter, and so light-protectivemechanisms had to be evolved to provide normal functioning under brighter illumination. Since the insect photoreceptor responds to light by depolarization, a fatal decrease in the resting potential down to zero might be caused by a strong light

178

F. G . GRIBAKIN

(e.g., direct sunlight), and again a need for protection from excessive light has arisen. At least two kinds of light-protection mechanisms are known at present: first, those diminishing the relative absorption of the rhabdomere, that is, optical mechanisms of sensitivity control; and, second, those controlling the transduction process electrically and chemically somewhere between the rhabdomere and the cell plasma membrane. In this respect vertebrate photoreceptors responding by hyperpolarization appear to have an advantage, since in the brightest environment all the sodium channels are closed, the membrane potential increases to obtain a steady level (light level), and the cell keeps all its energetic potency although its response to light ceases. Rhodopsin is a membrane protein, and in order for absolute sensitivity to evolve, the photoreceptor membrane had to increase in area. This should cause an inevitable increase in ionic leakage and consequently an increase in noise level. Thus it was of vital importance for natural selection to prevent generation of the receptor signal from being triggered by dark- or light-evoked sodium permeability changes in the photoreceptor membrane itself. An intracellular transmitter appears to have resolved this problem, so that no receptor potential can be generated by the cell plasma membrane until excited by the direct action of the transmitter. The nature of the transmitter is still unknown. The compound eye can be considered an insect body compartment, and the ionic composition of its extracellular medium is seemingly regulated by the activity of the photoreceptors themselves. As in vertebrates, there exists noticable dark current in insect photoreceptors, and total dark current in the eye can reach several microamperes. The clear spatial regularity of the insect eye allows one to consider this organ a ‘‘biological crystal, ” and workers investigating its integral properties can hope to obtain information on the properties of its constituents, that is, the photoreceptors and accessory cells. The crystallinity of the insect eye gives an advantage to a worker in neuromorphology and neurophysiology, and it is hoped that several general problems can be better understood using this model, for instance, principles of compartmentalizationof inorganic ions, mechanisms controlling the extracellular ionic composition of nerve tissue, antidromic axonal transport connected with direct (in this case, dark) current, and so on.

REFERENCES Abrahamson, E. W., and Wiesenfeld, J. R. (1972). Handb. Sens. Physiol. 7 , Part 1, 69-121. AUtNm, H . , and Kolb, G. (1968). 2. Vergl. Physiol. 60, 450-477. AutNm, H . , and Tschamtke, H. (1962). Z. Vergl. Physiol. 45, 695-710. Bader. C. D . , Baumann, F., and Bertrand, D. (1976). J . Gen. Physiol. 67, 475-491.

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

179

Baumann, C. (1972). Handb. Sens. Physiol. 7, Part 1, 395-416. Baumann, F. (1968). J. Gen. Physiol. 52, 855-875. Baumann, F. (1975). In “The Compound Eye and Vision of Insects” (G. A. Homdge, ed.), pp. 53-74. Oxford Univ. Press (Clarendon), London and New York. Baumann, F., and Hadjilazaro, B. (1972). Vision Res. 12, 17-31. Behrens, M., and Krebs, W. (1976). J. Comp. Physiol. 107, 77-96. Bernard, G. D. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 78-97. Springer-Verlag, Berlin and New York. Bernhard, C. D., and Ottoson, D. (1964). J. Gen. Physiol. 47, 465-478. Bemdge, M. J., and Prince, W.T. (1972). Adv. Insect Physiol. 9, 1-49. Bertrand, D., Perrelet, A., and Baumann, F. (1972). J. Physiol. (Paris) 65, 102A. Bikle, D., Tinley, L. G., and Porter, K. (1966). Protoplasma 61, 322-345. Blasie, J. K. (1972). Biophys. J. 12, 191-209. Borle, A. B. (1973). Fed. Proc., Fed. Am. SOC.Exp. Biol. 32, 1944-1950. Borsellino, A,, and Fuortes, M. G. F. (1968). J. Physiol. (London) 196, 507-539. Boschek, C. B., and Hamdod, K. (1976). 2. Naturforsch., Teil C 31, 763. Briggs, M. H. (1961). Nature (London) 192, 874-875. Brown, A. M.,Baur, P. S., and Tuley, F. H. (1975). Science 188, 157-160. Brown, H. M. (1976). J. Gen. Physiol. 68, 281-296. Brown, H. M., and Cornwall, M. C. (1975a). J . Physiol. (London)248, 555-578. Brown, H. M., and Cornwall, M. C. (1975b). J. Physiol. (London)248, 579-593. Brown, H. M., and Ottoson, D. (1976). J. Physiol. (London)257, 355-378. Brown, J. E., and Blinks, J. R. (1974). J. Gen. Physiol. 64, 643-665. Brown, P. K. (1972). Nature (London) New Biol. 236, 35-38. Brown, P. K., and White, R. H. (1972). J . Gen. Physiol. 59, 401-414. Bmckmoser, P. (1968). 2. Vergl. Physiol. 59, 187. Bumel, M., Mahler, H. R., and Moore, W. J. (1970). J. Neurochem. 17, 1493-1499. Burovina, I. V., Gribakin, F. G., Pogorelov, A. G., Petrosyan, A. M., Pivovarova, N. B., and Polyanovskii, A. D. (1978). J. Comp. Physiol. 127, 245-253. Burtt, E. T., and Catton, W. T. (1964a). J. Insect Physiol. 10, 689-710. Burtt, E. T., and Catton, W. T. (1964b). J. Insect Physiol. 10, 865-886. Butler, R. (1971). Z . Vergl. Physiol. 72, 67-80. Butler, R., and Horridge, G. A. (1973). J . Comp. Physiol. 83, 263-278. Bygrave, F. L. (1978). Biol. Rev. 53, 43-70. Carafoli, E., Tiozzo, R., Lugli, G., Crovetti, F., and Kratzing, C. (1974). J. Mol. Cell. Cardiol. 6, 361-371. Cherkasov, A. D., Vishnevskaya, T. M., and Mazokhin-Porshnyakov, G. A. (1976). In “Infomation Processing in Visual Systems,” (V. D. Glezer ed.), pp. 41-43. Nauka, Leningrad. Cone, R. A. (1972). Nature (London), New Biol. 236, 39-43. Cone, R. A,, and Brown, P. K. (1969). Nature (London) 221, 818-820. Cosens, D. J. (1967). J. Insect Physiol. 13, 1373-1386. Daeman, F. J. M. (1973). Biochim. Biophys. Acta 300, 255-288. Dartnall, H. J. A. (1953). Br. Med. Bull. 9, 24-30. Dartnall, H. J. A. (1972). Handb. Sens. Physiol. 7, Part 1, 122-145. Dartnall, H. J. A., and Lythgoe, J. N. (1965). Vision Res. 5 , 81-100. Dodge, F. A., Knight, B. W.,and Toyoda, J. (1968). Science 160, 88-90. Dowling, J. E., and Hubbard, R. (1963). Nature (London) 199, 972-975. Eakin, R. M. (1965). Cold Spring Harbor Symp. Quant. B i d . 30, 363-370. Eakin, R. M. (1972). Handb. Sens. Physiol. 7, Part 1, 626-684. Eakin, R. M., and Brandenburger, J. L. (1975). Am. 2001. 15, 851-863.

180

F. G. GRIBAKIN

Eckert, M. (1968). Zool. Jahrb., Abt. Allg. Zool. Physiol. Tiere 74, 102. Edidin, M. (1974). Annu. Rev. Biophys. Bioeng. 3, 179-204. Egner, 0. (1971). Cytobiologie 4, 262-292. Eguchi, E. (1971). 2. Vergl. Physiol. 71, 201-218. Eguchi, E., and Waterman, T. H. (1966). In “The Functional Organization of the Compound Eye” (C. G. Bemhard, ed.), pp. 105-124. Pergamon, Oxford. Eguchi, E., and Waterman, T. H. (1967). 2. Zellforsch. Mikrosk. Anat. 79, 209-229. Eguchi, E., and Waterman, T. H. (1968). Z . Zellforsch. Mikrosk. Anat. 84, 87-101. Eguchi, E., and Waterman, T. H. (1976). Cell Tissue Res. 169, 419-434. Eguchi, E., Waterman, T. H., and Akiyama, J. (1973). J. Gen. Physiol. 62, 355-374. Enoch, J. M. (1963). J. Opr. SOC. Am. 53, 71-85. Falk, G., and Fatt, P. (1972). Handb. Sens. Physiol. 7 , Part 1, 200-244. Femandez, H., and Nickel, E. (1976). J. Cell Biol. 69, 721-732. Finean, J. B., and Engstrom, A. (1967). “Biological Ultrastructure,” 2nd ed. Academic Press, New York. Franceschini, N. (1972). In “Information Processing in the Visual Systems of Arthropods” (R. Wehner, ed.), pp. 75-82. Springer-Verlag, Berlin and New York. Frantsevich, L. I., Zolotov, V. V., Gribakin, F. G., Polyanovskii, A. D., Govardovskii, V. I., and Zueva, L. V. (1976). Dokl. Akad. Nauk SSSR 226, 133-736. Frantsevich, L., Govardovskii, V., Gribakin, F., Nikolayev, G., Pichka, V., Polyanovskii, A., Shevchenko, V., and Zolotov, V. (1977). J. Comp. Physiol. 121, 253-271. Fulpius, B., and Baumann, F. (1969). J. Gen. Physiol. 53, 541-561. Fuortes, M. G. F., and O’Bryan, P. M. (1972a). Handb. Sens. Physiol. 7 , Part 2, 279-320. Fuortes, M. G. F., and O’Bryan, P. M. (1972b). Handb. Sens. Physiol. 7 , Part 2, 321-338. Fuortes, M. G. F., and Yeandle, S. (1964). J. Gen. Physiol. 47, 443-4453, Gogala, M. (1967). Z . Vergl. Physiol. 57, 232-243. Gogala, M., and Michieli, S. (1965). Naturwissenschafien 9, 217-218. Gogala, M., Hamdorf, K., and Schwemer, J. (1970). Z . Vergl. Physiol. 70, 410-413. Goldsmith, T. H. (1958). Proc. Natl. A c d . Sci. U.S.A. 44, 123-126. Goldsmith, T. H. (1965). J . Gen. Physiol. 49, 265-287. Goldsmith, T. H. (1972). Handb. Sens. Physiol. 7, Part 1, 685-719. Goldsmith, T. H. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 392-409. Springer-Verlag, Berlin and New York. Goldsmith, T. H., and Bernard, G. D. (1974). In “Physiology of Insecta” (M. Rockstein, ed.), 2nd ed., Vol. 2, pp. 165-272. Academic Press, New York. Goldsmith, T.H.,and Bruno, M. S. (1973). In “Biochemistry and Physiology of Visual Pigments” (H. Langer, ed.), pp. 147-153. Springer-Verlag, Berlin and New York. Goldsmith, T. H., and Femandez, H. R. (1966). In “The Functional Organization of the Compound Eye” (C. G. Bemhard, ed.), pp. 125-143. Pergamon, Oxford. Goldsmith, T. H., and Warner, L. (1964). J . Gen. Physiol. 47, 433-441. Goldsmith, T. H., and Wehner, R. (1975). Biol. Bull. (Woods Hole, Mass.) 149, 427-428. Goldsmith, T. H., and Wehner, R. (1977). J. Gen Physiol. 70, 453-490. Goldsmith, T. H., Barker, R. J., and Cohen, C. F. (1964). Science 146, 65-67. Gribakin, F. G. (1967a). Zh. Evol. Biokhim. Fiziol. 3, 66-73. Gribakin, F. G. (1967b). Tsitologiya 9, 1276-1280. Gribakin, F. G. (1969a). Tsitologiya 11, 308-314. Gribakin, F. G. (1969b). Nature (London) 223, 639-641. Gribakin, F. G. (1969~).Tr. Vses. Entomol. Obsch. 53, 238-273. Gribakin, F. G. (1972). Vision Res. 12, 1225-1230.

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

181

Gribakin, F. G. (1973). Nature (London) 246, 357-358. Gribakin, F. G. (1975). In “The Compound Eye and Vision of Insects” (G. A. Homdge, ed.), pp. 154176. Oxford Univ. Press (Clarendon), London and New York. Gribakin, F. G. (1979). Dokl. Akad. Nauk SSSR (submitted for publication). Gribakin, F. G., and Govardovskii, V. I. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 215-236. Springer-Verlag, Berlin and New York. Gribakin, F. G., and Petrosyan, A. M. (1979). J. Comp. Physiol. (submitted for publication). Gribakin, F. G., Pogorelov, A. G., and Burovina, I. V. (1976). Tsitologiya 18, 1176-1179. Gribakin, F. G., Petrosyan, A. M., Burovina, I. V., Pogorelov, A. G., and Leont’yev, V. G. (1977). In “The Sensory Reception Mechanisms,” (0. B. Ilyinskii, ed.), pp. 37-42. Nauka, Leningrad. Grundler, 0. J. (1974). Cytobiologie 9, 203-220. Guzzo, A. V., and Pool, G. L. (1968). Science 159, 312-314. Hagins, W. A,, and Liebman, P. A. (1963). Abstr. Biophys. SOC. 7th Annu. Meet. MEG. p. OOO. Hamdorf, K., and Razmjoo, S. (1977). Biophys. Struct. Mech. 3, 163-170. Hamdorf, K., and Schwemer, J. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 263-289. Springer-Verlag, Berlin and New York. Hamdorf, K., Schwemer, J., and Tauber, U. (1968). Z. Vergl. Physiol. 60, 375-415. Hamdorf, K., Schwemer, J., and Gogala, M. (1971). Nature (London) 231, 458-459. Hays, D., and Goldsmith, T. H. (1969). Z. Vergl. Physiol. 65, 218-232. Hecht, S., Shlaer, S., and Pirenne, M. H. (1942). J . Gen. Physiol. 25, 819-840. Heisenberg, M. (1971). J. Exp. Biol. 55, 85-100. Hillman, P., Keen, M. E., and Winterhager, J. W. (1977). Biophys. Strucr. Mech. 3, 183-190. Hochstein, S., Minke, B., and Hillman, P. (1973). J. Gen. Physiol. 62, 105-128. Hoglund, G. (1966). Acta Physiol. Scand. 69, Suppl. 282. Homdge, G. A. (1975). In “The Compound Eye and Vision of Insects” (G. A. Homdge, ed.), pp. 255-298. Oxford Univ. Press (Clarendon), London and New York. Homdge, G. A., and Barnard, P. B. T. (1965). Q. J. Microsc. Sci. [N.S.] 106, 131-135. Hubbard, R., and St. George, R. C. C. (1958). J. Gen. Physiol. 41, 501-528. Hubbard, R., and Wald, G. (1952). J. Gen. Physiol. 36, 269-315. Jan, L. Y., and Revel, J. P. (1974). J. Cell Biol. 62, 257-273. Kaplan, E., and Barlow, R. B. (1976). Vision Res. 16, 745-741. Kirschfeld, K. (1969). In “Processing of Optical Data by Organisms and Machines” (W. Reichardt, ed.), pp. 116-136. Academic Press, New York. Kirschfeld, K. (1974). Z. Narurforsch. Teil C 29, 592-596. Kirschfeld, K., and Franceschini, N. (1969). Kybernefik 6, 13-22. Kirschfeld, K., Franceschini, N., and Minke, B. (1977). Nature (London) 269, 386-390. Kirschfeld, K., and Snyder, A. W. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 56-77. Springer-Verlag, Berlin and New York. Kolb, G., and Autrum, H. (1972). J. Comp. Physiol. 77, 126-140. Kolb, G., and Autrum, H. (1974). J. Comp. Physiol. 94, 1-6. Langer, H. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 429-455. Springer-Verlag, Berlin and New York. Laughlin, S. B., Menzel, R., and Snyder, A. W. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 237-259. Springer-Verlag, Berlin and New York. Levinson, J. Z. (1972). Handb. Sens. Physiology 7 , Part 2, 339-356. Liebman, P. A. (1962). Biophys. J. 2 , 161-178. Liebman, P. A. (1972). Handb. Sens. Physiol. 7 , Part 1 , 481-528. Liebman, P. A., and Entine, G. (1968). Vision Res. 8, 761-775.

182

F. G. GRIBAKIN

Liebman, P. A,, and Entine, G. (1974). Science 185, 457-459. Liebman, P. A., Kaplan, M. W.,Jagger, W.S., and Bargoot, F. G. (1974). Nature (London) 251, 31-36. Lisman, J. E., and Brown, J. E. (1972). 1. Gen. Physiol. 59, 701-719. Lisman, J . E., and Brown, J. E. (1975). J. Gen. Physiol. 66, 489-506. Martinez, J. M., and Srebro, R. (1976). J. Physiol. (London)261, 535-562. Mason, W. T., Fager, R. S., and Abrahamson, E. W . (1974). Nature (London) 247, 188-191. Mazokhin-Porshnyakov, G. A. (1969). “Insect Vision. ” Plenum, New York. Menzel, R. (1975a). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 372-387. Springer-Verlag, Berlin and New York. Menzel, R. (197%). In “The Compound Eye and Vision of Insects” (G. A. Homdge, ed.), pp. 121-153. Oxford Univ. Press (Clarendon), London and New York. Menzel, R., and Knauth, R. (1973). J. Comp. Physiol. 86, 125-138. Menzel, R., and Lange, G. (1971). Z . Natu~orsch.Teil B 26, 357-359. Menzel, R., and Snyder, A. W .(1974). J. Comp. Physiol. 88, 247-270. Menzel, R., and Snyder, A. W. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 1-16. Springer-Verlag, Berlin and New York. Meyer-Rochow, V. B. (1974). J. Insect Physiol. 20, 573-589. Millecchia, R., and Mauro, A. (1969). J. Gen. Physiol. 54, 331-351. Miller, W. H. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 415-428. Springer-Verlag, Berlin and New York. Miller, W. H., and Cowthon, D. F. (1974). Invest. Ophrhalmol. 13, 401-405. Minke, B., Hochstein, S., and Hillman, P. (1973). J . Gen. Physiol. 62, 87-104. Minke, B., Wu, C.-F., and Pak, W. L. (1975). Nature (London) 258, 84-87. Moody, M. F., and Parks, J. R. (1961). 2. Vergl. Physiol. 44, 268-291. Morton, R. A. (1972). Handb. Sens. Physiol. 7, Part 1, 33-68. Morton, R. A., and Pin, G. A. J. (1969). Adv. Enzymol. 32, 97-171. Mote, M. I. (1970). J. Exp. Zool. 175, 149-158. Mote, M. I., and Goldsmith, T. H. (1971). Science 171, 1254-1255. Naka, K., and Eguchi, E. (1962). J. Gen. Physiol. 45, 663-680. Nemanic, P. (1975). Tissue & Cell 7, 453-468. Nickel, E., and Menzel, R. (1976). Cell Tissue Res. 175, 357-368. Nolte, J. E., and Brown, J. E. (1972). J. Gen. Physiol. 59, 186-200. Ottoson, D. (1964). J . Physiol. (London) 171, 109-118. Pak, W.L., Ostroy, S. E., Deland, M. C., and Wu, C.-F. (1976). Science 194, 956-959. Paulsen, R., and Schwemer, J. (1972). Biochim. Biophys. Acta 283, 520-529. Perrelet, A. (1970). 2. Zellforsch. Mikrosk. Anat. 108, 530-562. Perrelet, A , , and Bader C.-R. (1978). J. Ultrastr. Res. 63, 237-243. Petrosyan, A. M. (1977a). Candidate of Sciences Thesis, Sechenov Inst. Evol. Physiol. and Biochem., Leningrad. Petrosyan, A. M. (1977b). Tsitologiya 19, 284-287. Petrosyan, A. M., Gribakin, F. G., and Leont’yev, V. G. (1977). Zh. Evol. Biokhim. Fiziol. 13, 220-221. Polyanovskii, A. D. (1976). Tsitologiya 18, 1419-1427. Poo, M., and Cone, R. (1974). Nature (London) 247,437-441. Post, C. T., and Goldsmith, T. H. (1965). Biol. Bull. (Woods Hole, Mass.) 128, 473-487. Post, C. T., and Goldsmith, T. H. (1969). Ann. Entomol. Soc. Am. 62, 1497. Ranck, J. B. (1963). Exp. Neurol. 7, 144. Rohlich, P., and Toro, I. (1965). In “The Structure of the Eye” (J. W.Rohen, ed.), pp. 175-186. Schattauer, Stuttgart.

CELLULAR MECHANISMS OF INSECT PHOTORECEPTION

183

Rohlich, P., and Torok, L. (1962). Q. J. Microsc. Sci. [N.S.] 104, 543-548. Sakitt, B. (1972). J. Physiol. (London) 223, 131-150. Schmidt, W. J. (1938) Kolloid-2. 85, 137-148. Scholes, J. (1965). Cold Spring Harbor Symp. Quant. Biol. 30, 517-527. Schwemer, J. (1969). 2. Vergl. Physiol. 62, 121-152. Schwemer, J., and Paulsen, R. (1973). J . Comp. Physiol. 86, 215-229. Schwemer, J., Gogala, M., and Hamdorf, K. (1971). Z. Vergl. Physiol. 75, 174-188. Seitz, G. (1968). Z. Vergl. Physiol. 59, 205-231. Shaw, S. R. (1969a). Vision Res. 9, 999-1030. Shaw, S. R. (1969b). Vision Res. 9, 1031-1040. Shaw, S. R. (1972). J . Physiol. (London) 220, 145-175. Shaw, S. R. (1975). Nature (London) 255, 480-483. Shaw, S. R. (1977). J. Comp. Physiol. 113, 257-282. Sidman, R. (1957). J . Biophys. Biochem. Cytol. 3, 15-30. Singer, S. J., and Nicholson, G. L. (1972). Science 175, 720-731. Smola, U. (1976). I n “Neural Principles in Vision” (F. Zettler and R. Weiler, eds.), pp. 194-213. Springer-Verlag, Berlin and New York. Snitzer, E. (1961). J. Opt. SOC. Am. 51, 491-498. Snyder, A. W. (1973). J. Comp. Physiol. 83, 331-360. Snyder, A. W. (1975a). I n “Photoreceptor Optics” (A. W. Snyderand R. Menzel, eds.), pp. 38-55. Springer-Verlag, Berlin and New York. Snyder, A. W. (1975b). I n “The Compound Eye and Vision of Insects” (G. A. Homdge, ed.), pp. 179-225. Oxford Univ. Press (Clarendon), London and New York. Snyder, A. W., andHomdge, G. A. (1972). J. Comp. Physiol. 81, 1-8. Snyder, A. W., Menzel, R., and Laughlin, S. B. (1973). J. Comp. Physiol. 87, 99-135. Somjen, G. (1975). Annu. Rev. Physiol. 37, 163-190. Srebro, R., and Behbehani, M. (1974). J . Gen. Physiol. 64, 166-185. Stavenga, D. G. (1971). Proc. Int. Union Physiol. Sci., Int. Congr. 25th, 1971 Vol. 9, p. 532. Stavenga, D. G. (1974). J. Comp. Physiol. 91, 417-426. Stavenga, D. G. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 290-295. Springer-Verlag, Berlin and New York. Stavenga, D. G., and Kuiper, J. W. (1977). J. Comp. Physiol. 113, 55-72. Stavenga, D. G., Zantema, A,, and Kuiper, J. W. (1973). I n “Biochemistry and Physiology of Visual Pigments” (H. Langer, ed.), pp. 175-180. Springer-Verlag, Berlin and New York. Stavenga, D. G., Numan, J. A. J., Tinbergen, J., and Kuiper, J. W. (1977). J . Comp. Physiol. 113, 73-93. Stieve, H. (1964). Z. Vergl. Physiol. 47, 457-492. Stieve, H., Gaube, H., and Winterhager, J. (1976). Vision RPS. 16, 1159-1168. Swihart, S. L. (1972). J . Insect Physiol. 18, 1015. Taylor, A,, Mamelak, M., Reaven, E., and Maffly, R. (1973). Science 181, 347-350. Treherne, J. E., and Pichon, Y. (1972). Adv. Insect Physiol. 9, 257-313. Trujillo-Cenoz, O., and Melamed, J. (1966). J. Ultrastrucf. Res. 16, 395-398. Tsukahara, Y., and Horridge, G. A. (1977). J . Exp. Biol. 68, 137-149. Tsukahara, Y., Homdge, G. A., and Stavenga, D. G. (1977). J . Comp. Physiol. 114, 253-266. Tunstall, J., and Horridge, G. A. (1967). Z. Vergl. Physiol. 55, 167-182. Van?la, F. G., and Wiitanen, W. (1970). J. Gen. Physiol. 55, 336-358. Vinnikov, Ya.A. (1974). “Sensory Reception. Cytology, Molecular Mechanisms and Evolution. ” Springer-Verlag, Berlin and New York. Vishnevskaya, T. M., and Mazokhin-Porshnyakov, G. A. (1969). Biofizika 14, 151-157. Vishnevskaya, T. M., and Mazokhin-Porshnyakov, G. A. (1972). Biofizika 17, 635-641.

184

F. G. GRIBAKIN

Vishnevskaya, T. M., Cherkasov, A. D., Mazokhin-Porshnyakov, G. A., and Korzun, I. B. (1977). Biofizika 22, 323-327. Wachmann, E., Richter, S., and Schricker, B. (1973). Z. Morphol. Tiere 76, 109-128. Walcott, B. (1971). Z. Vergl. Physiol. 74, 17-25. Walcott, B. (1975). In “The Compound Eye and Vision of Insects” (G. A. Homdge, ed.), pp. 20-33. Oxford Univ. Press (Clarendon), London and New York. Wald, G. (1967). Nature (London) 215, 1131-1133. Wald, G., and Brown, P. K. (1953). J. Gen. Physiol. 37, 189-200. Wald, G., Brown, P. K., and Gibbons, I. R. (1962). Symp. SOC. Exp. Biol. 16, 32-57. Wasserman, G. S. (1973). Science 180, 268-275. Waterman, T. H. (1975). In “Photoreceptor Optics” (A. W. Snyder and R. Menzel, eds.), pp. 339-371. Springer-Verlag, Berlin and New York. Waterman, T. H., Fernandez, H., and Goldsmith, T. H. (1969). J. Gen. Physiol. 54, 415-432. Wehner, R. (1976). In “Neural Principles in Vision” (F. Zettler and R. Weiler, eds.), pp. 280-333. Springer-Verlag, Berlin and New York. Wehner, R., and Goldsmith, T. H. (1975). B i d . Bull. (Woods Hole, Mass.) 149, 450. Wehner, R., Bernard, G. D., and Geiger, E. (1975). J. Comp. Physiol. 104, 225-245. White, R. H. (1967). J. Exp. 2001.166, 405-425. White, R. H. (1968). J. Exp. 2001. 169, 261-278. White, R. H., and Lord, E. (1975). J . Gen. Physiol. 65, 583-598. Wikswo, M. A., and Novales, R. R. (1972). J. Ultrastruct. Res. 41, 189-201. Wolken, J. J., and Scheer, I. J. (1963). Exp. Eye Res. 2, 182-188. Wolken, J. J., Bowness, J. M., and Scheer, I. J. (1960). Biochim. Biophys. Acta 43, 531-537. Wulff, V. J., and Mueller, W. J. (1975). In “The Compound Eye and Vision of Insects” (G. A. Homdge, ed.), pp. 37-52. Oxford Univ. Press (Clarendon) London and New York. Wulff, V. J., Stieve, H., and Fahy, J. L. (1975). Vision Res. 15, 759-765. Yeandle, S. (1958). Am. J . Ophthalmol. 46, 82-87. Yeandle, S., and Spiegler, J. B. (1973). J. Gen. Physiol. 61, 552-571. Yoshizawa, T. (1972). Handb. Sens. Physiol. 7, Part 1, 146-179. Zettler, F., and Jarvilehto, M. (1971). Z. Vergl. Physiol. 75, 402-421.

NOTEADDEDIN PROOF Recent findings of P. G. Lillywhite strongly support the view that quantum bumps are unit responses of the locust photoreceptors to single photons [Lillywhite, P. G. (1977). J . Comp. Physiol. 122, 189-200; (1978). J. Comp. Physiol. 125, 13-27].