Light-Dependent Electrical Activity in Sea Urchin Tube Feet Cells.
The sea urchin has been used for many years as a model organism to identify key steps of animal fertilization and cellular events early in development (Briggs and Wessel, 2006). Much less, however, is known about the physiology and cell biology of adult urchins. In particular, the molecular events underlying the processes that enable sea urchins to sense and respond appropriately to their environment, for the most part, remain unknown. Echinoderms have relatively simple nervous systems consisting of a pentaradially symmetrical nerve ring from which radiate five radial nerves, which in turn innervate the numerous tube feet and spines (Cobb, 1985). How the sea urchin nervous system processes inputs from thousands of sensory cells that respond to light, touch, and chemical cues, to successfully navigate the environment (Reese, 1966), is unknown.
It has been known for many years that sea urchins can detect light (Millott, 1954), as can other echinoderms (Johnsen, 1994). It has also been suggested that Echinometra species and the purple sea urchin, Strongylocentrotus purpuratus, have a form of low-resolution vision (Blevins and Johnsen, 2004; Yerramilli and Johnsen, 2010), based on sea urchin phototaxis in the presence of dark shapes of varying angular width. In addition, definitive evidence of resolved vision has recently been demonstrated in Diadema africanum with the use of isoluminant visual stimuli (Kirwan et al., 2018). Light detection is believed to occur in sea urchins within photoreceptor cells in the terminal disc region of their motile tube feet, within spines, and across the test (Millot, 1954; Ullrich-Lifter et al., 2011). A typical behavioral response to bright illumination is seen in the negative phototaxis of S. purpuratus (Yoshida, 1966; Ullrich-Luter et al., 2011). How the light-dependent signals from thousands of disparate photoreceptor cells are integrated and processed in the simple nervous system of the sea urchin to initiate coordinated movement of hundreds of flexible tube feet and movable spines is unknown.
The sequencing of the S. purpuratus genome (Burke et al., 2006), the first of any echinoderm, has generated increased curiosity into the molecular components of sea urchin physiology. Analysis of the genome has revealed many genes that encode proteins that would be expected to be integral to light signal transduction (Burke et al., 2006), including six opsin genes (Raible et al., 2006) and orthologs of mammalian retinal transcription factors (Agca et al., 2011). Multiple opsin mRNAs have been detected in sea urchin tube feet (Raible et al., 2006; Ullrich-Luter et al., 2011), as well as opsin proteins; and the retinal transcription factor Pax6 has been found to be enriched in sea urchin tube feet discs (Agca et al., 2011; Lesser et al., 2011; Ullrich-Luter et al., 2011).
The initial step in the known light signal transduction pathways of animals is the absorption of photons, which isomerizes opsin-bound cofactors, allowing the activation of a G protein and either photoreceptor cell hyperpolarization or depolarization. To give some examples, in vertebrate rod and cone cells, opsin-mediated G protein activation leads to hyperpolarization (Takemoto and Cunnick, 1990) via the closure of cyclic nucleotide-gated (CNG) channels, whereas in some bivalves it leads to hyperpolarization via the opening of [K.sup.+] channels (Leung and Montell, 2017). Conversely, in mammalian intrinsically photosensitive retinal ganglion cells and arthropod photoreceptors, opsin-mediated G protein activation leads to depolarization (Minke and Selinger, 1996) via the opening of transient receptor potential (TRP) channels (Leung and Montell, 2017). In echinoderms, light flashes directed at optical cushion photoreceptors of the crown-of-thorns starfish (Acanthaster planci) elicit voltage changes (Petie et al., 2016). Although sea urchins clearly express opsin proteins and can detect light, the components and nature of the signaling events that transduce photon detection in electrical activity in the urchin nervous system are poorly understood.
In a previous study (Shah et al., 2018) we attempted to block the negative phototaxic response of S. purpuratus with the use of pharmacological inhibitors of enzymes and ion channels that are common to the light transduction pathways of other animals. However, most interventions blocked tube feet movement, which prevented us from disentangling the light detection steps from the movement response to light. In this study, therefore, we sought to isolate potential light detection steps that may occur in the tube feet discs from tube feet movement by using isolated tube feet disc cells. The electrical responses of individual tube feet disc cells in response to different levels of illumination were examined, as were the effects of altering extracellular ion concentrations on the currents.
Materials and Methods
Purple sea urchins, Strongylocentrotus purpuratus (Stimpson, 1857), about 60 mm in test diameter, from the Pacific Ocean off the coast of California (Marinus Scientific, Long Beach, CA) were maintained in aerated tanks at 13 [degrees]C containing artificial seawater (ASW) (Instant Ocean, That Fish Place, Lancaster, PA) made according to the manufacturer's instructions. The major ion concentrations of this ASW are (in mmol [L.sup.-1]): [Na.sup.+] (462), [K.sup.+] (9.4), [Mg.sup.2+] (52), [Ca.sup.2+] (9.4), [Cl.sup.-] (550), S[O.sup.2-.sub.4] (28), and [Li.sup.+] (20) (Atkinson and Bingman, 1997).
To isolate individual tube feet disc cells, we placed sea urchins in ASW with elevated [Mg.sup.2+] (73 mmol [L.sup.-1]), which facilitates the extension of their tube feet. Small dissecting scissors were used to cut off about 30-50 pigmented discs from the ends of extended tube feet. The tube feet discs were rinsed in ASW containing 2 mg m[L.sup.-1] glucose (glucose ASW) and then incubated for 90 minutes while gently rocking at room temperature in glucose ASW containing 2 mg m[L.sup.-1] collagenase. Partially dissociated tissue was then rinsed in glucose ASW and incubated for 60 minutes while gently rocking in glucose ASW containing 2.5 mg m[L.sup.-1] trypsin ethylenediaminetetraacetic acid. After a further wash in glucose ASW, tissue was pipetted through an 18-gauge needle 15-20 times and then through a 22-gauge needle 5-10 times. The dissociated cells were repeatedly rinsed in 0.22-[micro]m filtered glucose ASW until the solution became clear, and they were stored in ASW for 1-5 days at 13 [degrees]C, with most recordings performed on the same day as cell isolation. There was no correlation between the age of the cells and the type of currents observed.
Patch clamp recording
Cell-attached patch clamp recordings were performed on single isolated disc tube feet cells in suspension and visualized on an inverted microscope (Eclipse TE2000, Nikon, Minato, Tokyo) with a x40 objective. Currents were recorded using an Axopatch IB amplifier (Axon Instruments, San Jose, CA) coupled to a CV-3 headstage (Axon Instruments), using thick-walled (1.5-mm outer diameter, 0.86 inner diameter) borosilicate pipettes with resistances of 2-3 M[ohm]. Successfully sealed cells had membrane resistances in excess of 1 G[ohm]. Both the extracellular bath solution and the pipette solution consisted of ASW (see Cell isolation), with ion concentrations modified where stated. Data were acquired (Labscribe 2, iWorx, Dover, NH) at gains of 1-50 mV p[A.sup.-1] (based on current magnitudes), sampled at 10-50 kHz, and filtered at 2 kHz. All recordings were made at room temperature. The pipette potential was held at +80 mV except for in the reversal potential recordings, in which, once currents were observed, a voltage ramp was applied from +80 mV until the observed current reversed. Pipette solutions for reversal potential measurements comprised normal ASW or ASW with the indicated elevated concentrations of either NaCl, KC1, K-gluconate, or CaCL. Light modulation of currents was via a white light LED flashlight (model TN4A, ThruNite, Shenzhen, China) positioned 215 mm from the patched cell.
Measurement of LED light properties
At each of the 3 light intensity settings of the LED flashlight, spectra over the range of 200-1100 nm were recorded from a 10-cm integrating sphere, using a fiber-coupled spectrograph (Spec-10 cooled CCD, Roper Scientific, Martinsried, Germany) with a 5-[micro]m slit and a diffraction grating of 150 lines per millimeter. Power measurements were made using a 1830-C meter (Newport Corporation, Irvine, CA) and an 818-UV detector (Newport Corporation) with a 1-[cm.sup.2] active area with meter wavelength settings of 400, 500, 565, and 635 nm.
Acquired data were exported from Labscribe 2 and converted to PClamp 10 (Molecular Devices, San Jose, CA) readable text files, using a custom program written in R version 3.1.2 (R Core Team, 2014). Error bars are the standard errors of the means. All t tests are two-tailed paired tests and assume non-equivalent variance.
Cell-attached patches were obtained on single tube feet disc cells in suspension (Fig. 1A). Under brightly lit conditions (about 800 mW [cm.sup.-2]), no current responses were observed; however, switching to dark conditions resulted in large, slowly rising currents after a delay of several seconds (Fig. 1B, C). Dark conditions consisted of the ambient light within the light-shielded electrophysiology rig in the absence of LED illumination (less than 1 mW [cm.sup.-2]). Two types of responses to the illumination protocol were observed. Currents designated Type I (64% of cells, n = 22) produced currents that plateaued following removal of illumination and that decayed upon re-illumination (Fig. 1B). The remaining 36% of cells elicited currents designated as Type II, characterized by currents induced by dark conditions that decayed spontaneously without any re-illumination (Fig. 1C). No individual cell exhibited both Type I and Type II currents in repeated testing, and out of a total of 97 cells recorded under multiple different conditions, only 3 cells failed to elicit any response.
The Type I current onset was characterized by two phases: a relatively fast sigmoidal phase and a very slow latter phase, whereas the current decay seemed to consist solely of a relatively fast phase (Fig. 1C). The majority of the current onsets and decays observed could not be adequately fitted with either sigmoidal or exponential functions; therefore, current onset and decay were quantified by measuring current half-time, defined as the time taken for the current to rise or decay to half of the maximum current value. No significant difference between the mean onset current half-time and the mean decay current half-time was detected (Student's t test, P = 0.97; n = 11 and n = 6 for current onset and decay, respectively; Fig. 1D).
To determine whether the electrical spikes generated when the light source was turned on or off were in some way responsible for the observed Type I or Type II currents, we recorded a patched cell while turning the light source off and then on, while obscuring all of the emitted light. Brief spikes but no sustained currents were observed (Fig. 1E). In addition, to determine the effect on the recording apparatus of switching the flashlight on and off, a patch pipette was patched onto the base of a petri dish coated with the inert material Sylgard (Dow, Midland, MI). Brief current spikes occurred when the flashlight was switched off and on in these control experiments, but no sustained currents were observed (Fig. 1F).
The spectral properties of the white LED light source were determined over the visible light range; major peaks were seen at 447 and 565 nm (Fig. 2A). Further recordings were made from tube feet disc cells at each of the three light intensity settings of the light source. Currents recorded from tube feet disc cells decreased with increasing irradiance (Fig. 2B), with a mean maximum observed current of 2231 [+ or -] 343 pA in the dark and effectively no currents at the highest irradiance level tested.
In the cell-attached patch clamp configuration, it is not possible to know the true cell membrane potential because there is no access to the intracellular environment of the cell; therefore, the membrane reversal potential cannot be estimated. However, the pipette holding potential is known, and changes in specific pipette solution ion concentrations should alter the pipette reversal potential if these specific ions contribute to the generated currents. A voltage ramp, ranging from +80 to +110 mV, was applied to a Type I cell in our standard ASW, and the pipette reversal potential was determined to be +94.2 [+ or -] 1.3 mV, n = 9 (Fig. 3A). Increasing the concentration of NaCl in the pipette ASW solution by 20% and by 50% led to significant decreases in pipette reversal potentials (ANOVA, P = 0.00004; n = 3 for control, n = 5 for 20% and 50% additional NaCl; Fig. 3B). Similarly, increasing the concentration of CaCL in the pipette ASW solution by 20% and by 50% also led to significant decreases in the pipette reversal potentials (ANOVA, P = 4 x [10.sup.-8]; n = 3 for control, n = 5 for 20% and 50% additional Ca[Cl.sub.2]; Fig. 3C). Increasing the concentration of [K.sup.+] in the pipette ASW solution by 20% or by 50% with either KCl or [K.sup.+]-gluconate also significantly decreased the pipette reversal potentials (ANOVA, P = 3 x [10.sup.-9]; n = 3 for control, n = 4 for 20% additional KCl, n = 5 for 50% additional KCl, n = 3 for 20% additional [K.sup.+]-gluconate, and n = 4 for 50% additional [K.sup.+]-gluconate; Fig. 3D).
Closer examination of the off-currents revealed them to be composed of the summation of multiple, smaller events. After removal of illumination there was a delay of several seconds and then abrupt stepwise increase in current (Fig. 4A, B). Each stepwise current increase consisted of a very rapid (on the order of 1-4 ms) current increase, followed by a slower current decay, which decayed to a new baseline current greater than before the rapid event. The decay of these "miniature" events could be well fitted with a single exponential curve with a mean time constant of 20.8 [+ or -] 3.5 ms (n = 20; Fig. 4C). The current magnitude of the exponentially decaying current events ranged from about 1 pA to several hundred picoamperes, and the summation of many of these miniature events seemed to account completely for the large dark-activated currents seen.
In this study we developed a protocol for the isolation of, and electrical recording from, individual sea urchin tube feet disc cells. We functionally identified two different types of tube feet disc cells; both cell types generated currents upon the removal of illumination. However, Type I cell currents were sustained until re-illumination, whereas Type II cell currents decayed prior to re-illumination. It may be that the two functional cell types observed represent two functionally different types of cells or, alternatively, that they may represent the same types of cells but in different stages of health.
The increase in electrical activity seen in both cell types upon removal of illumination has some parallels to that seen in direct recordings from the radial nerve of the sea urchin Diadema setosum in response to illumination (Takahashi, 1964). In these experiments, electrical activity of the nerve was detected upon changing the illumination level both from light to dark and from dark to light; however, a greater duration of activity was seen with the light-to-dark transition compared to the dark-to-light transition. Interestingly, as with our data, the delay between the switching of the light source and the electrical responses observed in the earlier study (Takahashi, 1964) were on the order of several seconds. Given this relatively long delay, it seems likely that the removal of illumination is not directly altering ion channel activity, but it may be activating multiple biochemical steps prior to the observed current activation.
From these cell-attached patch clamp experiments we are unable to determine the relative ion permeabilities of the current. However, the shifts in pipette reversal potential when altering the pipette concentrations of NaCl, Ca[Cl.sub.2], KC1, and [K.sup.+]-gluconate indicate that [Na.sup.+], [Ca.sup.2+], and [K.sup.+] are all involved in generating these currents. This indicates an ion-conducting pathway of relatively low selectivity, qualitatively similar to the low selectivity of both CNG and TRP channels found in the visual signal transduction pathways of many other animals.
The activation of a current upon the removal of illumination (known as an off-current) seen here in the urchin tube feet disc cells has some similarities with the vertebrate light detection system found in rod and cone cells, in which the absorption of light leads to an enzyme cascade that results in the turning off of a constitutive dark current (Takemoto and Cunnick, 1990). Similarly, off-currents have been observed in scallop visual cells (Wilkens, 2008). These exampies contrast with the electrical responses of arthropod photoreceptor cells, in which illumination leads to an enzyme cascade that activates photoreceptor cell currents (e.g., in Drosophila via the opening of TRP channels; Minke and Selinger, 1996); and, conversely, currents are turned off by channel closure upon removal of illumination. In both the vertebrate rod and cone and the scallop systems, current inhibition is due to photoreceptor hyperpolarization, albeit via different mechanisms. In vertebrates, closure of cation-permeable CNG channels hyperpolarizes rods and cones in response to light, whereas in scallops, the opening of CNG [K.sup.+] channels underlies the hyperpolarizing response. Given the permeability data of the sea urchin tube feet cells, it would seem that it is not solely a [K.sup.+] conductance that is responsible for the current inhibition seen in response to light.
The observed dark-induced currents were seen to be composed of discrete, smaller events that activated rapidly and decayed exponentially. The variable amplitude and the exponential decay of these events indicate that they are not due to the opening of individual ion channels, which would instead display discrete stepwise increases and decreases in current (Hamill et al., 1981). Superficially, the events look similar to postsynaptic miniature end plate currents that can be observed in postsynaptic neurons in response to a presynaptic vesicular releaser of a neurotransmitter. However, the miniature events observed could not be postsynaptic responses because they were recorded from individual, isolated cells. However, a similar current time course would be expected if vesicles within the tube feet disc cells were fusing to the membrane within the pipette patch, with the onset of the current representing the opening of the fusion pore between the vesicle and the pipette solution, and the exponential decay of current representing the depletion of ions within the vesicle into the patch pipette. Indeed, similar current time courses have been observed previously in studies that directly measured vesicular fusion by using the patch clamp technique (Breckenridge and Aimers, 1987; Calvo-Gallardo et al., 2015). Numerous vesicles of differing sizes have been seen in electron micrographs of urchin photoreceptor cells from tube feet discs (Ullrich-Luter 2011), and vesicles of differing size would be expected to generate fusion currents of variable amplitudes, as were seen in this study. The function of vesicle fusion in sea urchin tube feet cells has been hypothesized to aid in membrane turnover for the recycling of membrane components. Alternatively, because sea urchin tube feet disc photoreceptor cells in vivo connect directly to tube foot nerves (Ullrich-Luter et al., 2011), the miniature currents we observed may represent synaptic vesicular release from sea urchin tube feet disc cells as part of the expected onward signaling from photoreceptor cells to the rest of the sea urchin nervous system.
In conclusion, we have developed a protocol to isolate and record electrical activity from individual sea urchin tube feet disc cells. Surprisingly, electrical activity of the sea urchin tube feet disc cells was inhibited upon illumination and was stimulated in the absence of light, and it resembled vesicular fusion occurring within the recording pipette.
This work was supported by funding from Franklin and Marshall College (Faculty Research Grant to CS, a Hackman Summer Scholarship to LJM, and an Independent Study Award to MAS) and the University of the South (Kresge and Conduff funds to CS). Many thanks to Dr. Eugenii U. Donev (University of the South, Department of Physics) for aiding in the measurement of the LED light spectral and power properties.
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LAUREN J. MARCONI (1,*), AVERY STIVALE (1), MUNEEB A. SHAH (1), AND CHRIS SHELLEY (1,2,[dagger])
(1) Department of Biology, Franklin and Marshall College, Lancaster, Pennsylvania 17604; and (2) Department of Biology, University of the South, 735 University Avenue, Sewanee, Tennessee 37383
Received 18 September 2017; Accepted 15 November 2018; Published online 25 February 2019.
(*) Present address: Department of Pharmacology, University of Maryland School of Medicine, 655 West Baltimore Street, Baltimore, Maryland 21201.
([dagger]) To whom correspondence should be addressed. E-mail: email@example.com.
Abbreviations: ASW. artificial seawater; CNG. cyclic nucleotide-gated; TRP, transient receptor potential.
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|Author:||Marconi, Lauren J.; Stivale, Avery; Shah, Muneeb A.; Shelley, Chris|
|Publication:||The Biological Bulletin|
|Date:||Apr 1, 2019|
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