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1Department of Biological Sciences, Rutgers University; and 2Department of Mathematical Sciences, New Jersey Institute of Technology, Newark, New Jersey
Submitted 12 April 2006; accepted in final form 26 June 2006
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ABSTRACT |
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INTRODUCTION |
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; Desai et al. 1999; Franklin et al. 1992; Hong and Lnenicka 1995; Li et al. 1996; Linsdell and Moody 1994; Turrigiano et al. 1994) and in neuronal networks (Galante et al. 2001; Luther et al. 2003; Turrigiano and Nelson 2004). This homeostatic plasticity allows neuronal and network activity to remain within functional limits during their normal operation and in response to perturbations or injury. Regulation of neuronal excitability and intrinsic properties can be induced by electrical stimulation (Cudmore and Turrigiano 2004; Franklin et al. 1992; Garcia et al. 1994; Golowasch et al. 1999a; Li et al. 1996; Turrigiano et al. 1994), ionic conductance perturbations (Desai et al. 1999; Linsdell and Moody 1994), development and growth (Spitzer et al. 2002), and trauma (Darlington et al. 2002) and may play a role in memory formation (Daoudal and Debanne 2003; Marder et al. 1996; Zhang and Linden 2003).
The rhythmical activity pattern of the pyloric network in the stomatogastric ganglion (STG) of crustaceans is conditional on neuromodulatory release from neurons located in central ganglia. When these inputs are permanently removed, activity ceases but a stable new activity pattern spontaneously develops within hours to days (Golowasch et al. 1999b; Luther et al. 2003; Thoby-Brisson and Simmers 1998). It has been suggested that this recovery of rhythmic activity occurs via up- and downregulation of voltage-dependent ionic conductances and the consequent acquisition of oscillatory properties by some of the network components (Golowasch et al. 1999b; Mizrahi et al. 2001; Thoby-Brisson and Simmers 2002). Indeed, this recovery correlates with some ionic conductance changes (Mizrahi et al. 2001; Thoby-Brisson and Simmers 2002). Neuromodulators may also have suppressive trophic effects on the excitability of STG neurons in the intact network, suppression that may be released after removal of neuromodulatory input by decentralization (Le Feuvre et al. 1999; Thoby-Brisson and Simmers 1998, 2000).
Neuronal intrinsic excitability is affected by patterned electrical activity in isolated STG neurons in culture (Turrigiano et al. 1994) and in neurons in situ (Golowasch et al. 1999a). Excitability changes can also occur spontaneously both in isolated STG neurons in culture in response to cell dissociation (Turrigiano et al. 1995) or in the intact ganglion in response to decentralization (Golowasch et al. 1999b; Mizrahi et al. 2001; Thoby-Brisson and Simmers 2002). To understand how network activity evolves in response to persistent perturbations (such as the removal of central inputs essential for the generation of the activity), it is essential to understand the dynamics and plasticity of the voltage-dependent ionic currents of its component neurons. In cultured STG neurons, spontaneous activity changes have been correlated with changes in a TEA-sensitive K+ current and in various inward currents (Turrigiano et al. 1995). However, the conductance changes induced by prolonged rhythmic stimulation (Turrigiano et al. 1994) have not been identified.
Here we study the ionic mechanisms involved in spontaneous and activity-induced recovery of oscillatory activity in adult dissociated crab STG neurons. We find the same ionic currents to be modified in both cases, although probably via different signaling pathways. Dynamic-clamp experiments confirm that the ionic currents modified are sufficient to produce the observed activity changes. Surprisingly, during the first 10 days after dissociation, neurons from the pyloric and gastric networks cannot be distinguished on the basis of neuronal activity, ionic conductance changes, or response to stimulation.
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METHODS |
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Adult male crabs Cancer borealis (carapace length >15 cm) were obtained from local fish markets (Newark, NJ) and maintained in saltwater aquaria at 12–14°C. The following solution compositions were used (concentrations all in mM): standard Cancer saline solution (440.0 NaCl, 11.0 KCl, 13.0 CaCl2, 26.0 MgCl2, 5.0 Maleic acid, and 11.2 Trizma base, pH 7.4–7.5); salt supplement solution (743.7 NaCl, 16.4 KCl, 24.7 CaCl2, 50.2 MgCl2 and 10.0 HEPES, pH 7.4); 0 Ca2+/0 Mg2+ dissociation solution (440.0 NaCl, 11.0 KCl, and 10.0 HEPES, pH 7.4); barium saline solution (440.0 NaCl, 11.0 KCl, 12.9 BaCl2, 0.1 CaCl2, 26.0 MgCl2, 5.0 Maleic acid, and 11.2 Trizma base, pH 7.4–7.5). Mn2+ saline was identical to barium saline with Mn2+ substituting for Ba2+. All chemicals were obtained from Fisher Scientific (Fairlawn, NJ) unless otherwise indicated. The sodium channel blocker Tetrodotoxin, TTX (EMD, Biosciences) was used at 0.1 µM.
Cell dissociation
Adult STG neurons were cultured following a protocol similar to those used by Turrigiano and Marder (1993), Glowik et al. (1997), and Swensen and Marder (2000). Crabs were anesthetized by cooling during 15–30 min on ice. The foregut was removed, and the STG, with a portion of the nerves attached, was isolated as previously described (Selverston et al. 1976) in a sterile laminar flow hood. The dissected nerves and ganglia were rinsed four to five times in sterile Cancer saline containing 0.1 mg/ml gentamicin (MP Biomedicals, Aurora, OH). The ganglia were pinned down in sterile silicone elastomer (Sylgard)-lined Petri dishes, incubated in sterile 0 Ca2+/0 Mg2+ saline plus 2 mg/ml of the proteolytic enzyme Dispase (Gibco) for 6 h at room temperature, and then transferred to an incubator at 12°C overnight in the same solution. Individual somata were then removed from the ganglia by aspiration with glass micropipettes coated inside with goat serum (Invitrogen, Carlsbad, CA) and with fire-polished tips. Dissociated neurons were plated individually onto uncoated 35-mm plastic Nunclon culture dishes in sterile salt supplement solution diluted 1:1 with sterile Leibowitz L-15 medium (Invitrogen) and then placed in an incubator at 12–14°C for the duration of the culturing period. Saline was not replaced during this time.
Cells were discarded if they showed blebs protruding from the cell body and if they were not firmly attached to the substrate. Cells with primary neurite and cell bodies firmly attached to the substrate were found to be the healthiest and produced the most stable recordings. To verify that the proteolytic treatment did not unduly affect the cells during dissociation, we performed two separate tests. In one, we followed the enzymatic treatment as described in the preceding text and then tested the response of the entire pyloric network to neuromodulators. Figure 1A shows one example in which the pyloric activity [characterized by the alternating bursting of the lateral pyloric (LP), pyloric constrictor (PY), and pyloric dialator (PD) neurons] was recorded extracellularly from the pyloric network’s main motor nerve (the lateral ventricular nerve, lvn). Control before enzymatic treatment is shown in the top trace. After 24 h of proteolytic treatment (control—24 h, middle), a clear effect of the enzymatic treatment can be seen by comparing the top two traces. Immediately before the mechanical dissociation was performed (i.e., 24 h after the beginning of dispase treatment) 100 µM pilocarpine was applied. A response to the neuromodulator typical of this system (Nusbaum and Beenhakker 2002) can be observed. This is characterized by an increase in the number of spikes per burst and in the frequency of the pyloric rhythm. As a second test, we applied 1–10 µM pilocarpine to isolated STG neurons 1–5 days after enzymatic treatment and mechanical dissociation. Approximately 50% of the neurons responded by changing their activity pattern (Fig. 1B). Note that not all neurons in the intact STG respond to pilocarpine (Swensen and Marder 2001). Thus both of these tests confirm that neurons retain many of their biophysical properties through the proteolytic and mechanical dissociation procedure, as has been reported previously (Panchin et al. 1993; Turrigiano and Marder 1993).
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A subset of the experiments was performed with identified neurons of the STG. Because the gastric network is known to oscillate at a frequency several times slower than the pyloric network (Nusbaum and Beenhakker 2002), we hypothesized that gastric neurons, if they become oscillatory in dissociated culture, would oscillate at a lower frequency than dissociated pyloric neurons. Thus we aimed to distinguish only between neurons belonging to the pyloric network, the gastric network and other neurons not belonging to either of these networks. For this purpose, we followed the established protocol for cell identification (Selverston et al. 1976). A map of the STG neurons was drawn and neurons successively impaled with one electrode and identified. The tips of the theta filament electrodes used for recording (WPI, Sarasota, FL) were filled with either the dye Alexa Fluor 488 or Alexa Fluor 568 (Invitrogen) by dipping the back of the electrode in the dye solution for 1–2 min and backfilling the rest of the micropipette with 1 M KCl. Neurons identified as pyloric were filled by passing –10 nA for 10–30 s with one dye, and then neurons identified as gastric were filled similarly with the other dye. After this, the dissociation procedure was followed as described above.
Electrophysiological recordings
Single- or two-electrode voltage clamp (SEVC or TEVC) performed with an Axoclamp 2B amplifier (Molecular Devices, Union City, CA) was used to measure ionic currents. Data were digitized and then analyzed using the pClamp9.0 software (Molecular Devices). Recordings were obtained using Citrate-filled microelectrodes (4 M K-citrate +20 mM KCl). Current injection electrodes had resistances of 12–18 M
and voltage recording electrodes of 15–25 M. The preparation was grounded using an Ag/AgCl wire connected directly to the bath or via an agar bridge (4% agar in 0.6 M K2SO4 + 20 mM KCl). No differences were observed with or without the agar bridge. All experiments were carried out at room temperature (20–22°C) 
1 h after transferring the cells to the recording setup.
K+ currents were measured in standard Cancer saline +0.1 µM TTX and separated into two components: high-threshold, voltage-gated currents, IK, activated with 800-ms-long depolarizing membrane potential steps from a holding voltage of –40 mV, and the voltage-gated transient current, IA, that was activated with depolarizing steps from a holding voltage of –80 mV (Golowasch and Marder 1992; Graubard and Hartline 1991). In crabs, the high-threshold component is known to consist of two conductances, a delayed-rectifier, IKd, partially blocked by TEA and a Ca2+-dependent conductance, IK(Ca), that is completely blocked by 10–20 mM TEA and also indirectly by Cd2+, which blocks the underlying Ca2+ current (Golowasch and Marder 1992; Hurley and Graubard 1998). The high-threshold currents activated during the IA activation protocol were removed by subtracting the currents measured from a holding potential of –40 mV from those obtained from a holding voltage of –80 mV. The hyperpolarization-activated current, Ih, was measured using 4-s-long hyperpolarizing pulses from a holding potential of –40 mV. The Ca2+ current, ICa, was measured with depolarizing membrane potential steps from a holding voltage of –40 mV after blocking outward currents. For this, one electrode, filled with 4 M K-citrate +20 mM KCl, was used to record voltage, whereas the second, filled with 1 M TEA +1 M CsCl (12–25 M resistance) was used to inject current. Additionally, 20 or 100 mM TEA + 0.1 µM TTX was added to the standard Cancer saline bathing solution. Furthermore, in some experiments, we measured Ca tail currents at –80 mV induced with pulses to depolarized membrane potentials. The K+ equilibrium potential in these cells was found to be very close to –80 mV (not shown). At that voltage, ICa tails inactivate slowly, allowing us to determine ICa largely uncontaminated by K+ currents. To construct I-V plots, ICa was measured by averaging the current 20–30 ms after the onset of the voltage pulse (vertical arrow in Fig. 6B and C). Currents were leak subtracted using the p/n subtraction method included in the data-acquisition software, with the holding voltage during leak pulses at –40 mV and applying five subpulses of opposite polarity. For all voltages tested, these leak subtraction subpulses remain in the linear range of the current-voltage relationship. The current carried by Ba2+ through Ca2+ channels was also sometimes measured. For this, the barium saline described in the preceding text plus 20 mM TEA and 0.1 µM TTX was used.
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). In all cases, we also determined conductance and midpoint of voltage-dependent activation by fitting conductance versus voltage curves with a Boltzman equation (see equations in the following text). IKd and Ih were measured at steady state, whereas IA and ICa were measured at the peak (25–80 ms after pulse onset). Currents within a given experimental group were normalized by dividing all currents by the current value measured in control at 0 mV for all K+ currents (Figs. 6–8), at +10 mV for ICa (Figs. 6 and 7) and +20 mV for ICa tail (Fig. 6C). We chose these voltages to maximize the numbers of cells we could use for our analyses because not all cells could be voltage clamped to higher levels. The leak conductance was measured either as 1/Rin, where Rin is the input resistance measured with small hyperpolarizing current steps in current clamp, or as the slope of the I-V curve in the voltage range where no voltage-dependent currents appear to be activated (–40 to –60 mV).
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Capacitance was determined from the areas of the capacitive current transient resulting from 10-ms-long hyperpolarizing membrane potential steps from a holding voltage of –40 or –50 mV. In some neurons, we estimated the electrotonic length, L, by applying a low-amplitude hyperpolarizing current pulse around the resting potential of the cells. This was repeated 20 times, and the traces averaged to reduce noise. Double-exponential fits to the voltage change produced the membrane time constant, 
0, and an equalizing time constant, 1, which were used to determine L according to L = 
/
0/1 – 1 (Rall 1977).
Stimulation protocols
Neuronal activity was altered by applying 1-s-long hyperpolarizing current pulses at 0.33 Hz, driving neurons to a membrane potential of approximately –120 mV (peak response, see Fig. 4E). This protocol has been found to be effective in modifying the spontaneous activity of cultured lobster STG neurons before (Turrigiano et al. 1994). Current level was adjusted often during the stimulation period to maintain this level of hyperpolarization. Control ionic current measurements were recorded in voltage clamp immediately prior to the beginning. Currents were measured again after a 45- to 60-min stimulation period.
Analysis
SigmaStat (Aspire Software International, Leesburg, VA), Origin (OriginLab, Natick, MA), and CorelDraw (Corel) software packages were used for statistical and graphical analysis. ANOVA tests were performed using either a standard two-way ANOVA, the nonparametric Kruskal-Wallis ANOVA on ranks for nonnormally distributed populations, or a repeated-measures (RM) ANOVA. Results of statistical analysis were considered significant if the significance level P was below 
= 0.05. All error bars shown and the reported variability around the averages correspond to SDs.
Dynamic-clamp experiments
A NI PCI-6070-E board (National Instruments, Austin, TX) was used for current injection in dynamic clamp experiments. Data acquisition was performed using the Digidata board and pClamp software as described in the preceding text. The dynamic-clamp software was developed by Farzan Nadim and collaborators (available for download at http://stg.rutgers.edu/software.htm) in the LabWindows/CVI software environment (National Instruments, Austin, TX) on a Windows XP operating system. Ionic currents that showed an approximately average effect in response to prolonged rhythmic stimulation were recorded and fitted with Hodgkin and Huxley-type equations. We used the measured parameters that produced the best fit to reproduce ionic conductances that were then added to or subtracted from a neuron using dynamic-clamp (Sharp et al. 1993) adjusting only the maximum conductances to match the effects of rhythmic stimulation.
The equations we used to characterize each ionic current IX are
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mX and hX are the time constants with which the m and h gates, respectively, evolve in time toward their respective steady states mX
and hX. These steady states are governed each by two voltage-dependent parameters V1/2X and sX that are listed in Table 1. q is an exponent that takes value 1 if the ionic current exhibits voltage-dependent inactivation and value 0 if it does not.
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RESULTS |
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Our dissociation procedure typically yielded around 20–40% of the 25–26 STG neurons found (Kilman and Marder 1996) in each ganglion and, with careful suction, the soma and a relatively long segment of the major neurite could be removed. Figure 2, B–D, illustrates the morphology of three typical neurons with differing neurite lengths immediately after dissociation (days 0 and 1) and 6 days later. By the sixth day in culture, some neurons grew wide lamellipodia (Fig. 2B), others tended to grow one or more long processes with smaller lamellipodia extending from the ends (Fig. 2D), whereas others showed a combination of relatively wide lamellipodia and long processes (Fig. 2C). Any significant outgrowth originated almost exclusively from the neurite stump (Fig. 2, C and D) however short it may have been (see Fig. 2B). Only very small extensions were occasionally observed growing directly out of the soma (Fig. 2C, day 6). No obvious correlation was observed between the length of the original neurite and subsequent outgrowth morphology or electrical activity.
Neurons were classified according to the pattern of activity they expressed. Three types of electrical activity were readily distinguishable from the first day in culture when the cultured neurons were depolarized with low-amplitude current injection: silent, tonic, or oscillatory. Neurons that responded passively to all depolarizing and hyperpolarizing current injection levels were classified as silent (day 1 in Fig. 2, B and C).Tonic neurons were those that exhibited fast, transient depolarizations having a duty cycle 
0.2 (Fig. 2A1, top, and B, day 4) and that could be blocked with 0.1–1 µM TTX (Fig. 2A1, bottom). Oscillatory activity was defined as slow, low-amplitude depolarizations having a duty cycle >0.2 (Fig. 2A2, top and middle, also see day 6 in Fig. 2, B–D), resistant to TTX treatment (Fig. 2A2, middle) and sensitive to Ca2+ current blockers such as Mn2+ (Fig. 2A2, bottom) or Cd2+ (not shown). The oscillations recorded on days 4 and 6 of Fig. 2C are considered to be at a transition between tonic and oscillatory states and were classified as oscillatory. Duty cycle was defined as the duration of a depolarizing event (action potential or slow oscillation) relative to the period of an oscillation measured at 50% of the maximum oscillation amplitude.
None of the neurons recorded during the initial 10 days in culture expressed spontaneous activity without some depolarizing current, and all neurons were tested with depolarizing current steps of various amplitudes. On day 0, most neurons (81%) are silent, 14% of the neurons showed tonic firing of action potentials, and a very small percentage (<5%) of the nearly 350 neurons we recorded from expressed oscillatory activity (Fig. 3A).
Figure 3A shows that during days 0–3 in culture, the silent state of activity was dominant, but the proportion of oscillatory neurons steadily increased as time progressed. The proportion of tonic neurons began to increase at day 3, whereas the proportion of silent neurons steadily decreased. The proportion of silent neurons continued to decrease throughout the initial 7 days, whereas that of oscillatory neurons steadily increased (with the exception of day 4). Instead, the proportion of tonically firing neurons seemed to reach a plateau on day 4 (Fig. 3A). The rate of change of neuronal activity we observed was slower than that previously observed in cultured lobster neurons (Turrigiano et al. 1995). This may be due to the fact that we incubated our dissociated cells at 12°C instead of 20°C and/or to species differences.
To determine whether any of the activity changes correlate with STG neuron identity, we recorded from 37 additional neurons, classified into pyloric and gastric type (see METHODS). Of these, 25 produced oscillations as defined in the preceding text at days 2–8 in culture. The average oscillation frequency of the identified pyloric neurons was 3.65 ± 2.01 Hz (n = 14), whereas the average oscillation frequency of the identified gastric neurons was 2.95 ± 1.43 Hz (n = 11). Thus the average oscillation frequency of the gastric neurons was somewhat lower than that of the pyloric neurons but this difference was not statistically significant (P = 0.342, Student’s t-test).
Spontaneous conductance changes with time in culture
To understand the contributions of ionic currents to the activity changes described in the preceding text, we measured individual ionic currents at different times in culture and estimated their conductance. Figure 3, B and C, shows the evolution of five different ionic conductances over 10 days in cell culture. When data are available, we show conductance density changes over time rather than simply conductance changes to remove possible effects of growth. Only two of these showed statistically significant changes over that period (Fig. 3B) after normalizing by the capacitance of the cell: gCa, which increased by 159% (P = 0.002, n = 38), and the high-threshold outward current gK that includes both a delayed rectifier and a Ca2+-dependent K+ current (Golowasch and Marder 1992), which decreased by 54% (P = 0.047, n = 65, Fig. 3B). gleak increased by 79% (P < 0.001, n = 139, Fig. 2C) but when normalized by cell capacitance, the increase was reduced to only 15% and was not statistically significant (P = 0.777, n = 32). Thus the change in leak conductance can probably be explained simply by the growth of the cell, whereas both gCa and gK changes appear to be related to changes in neuronal activity as these changes correlate with the progression from silent to tonic to oscillatory activity (Fig. 3A). In contrast, gA and gh (Fig. 3C) increased by 30% between days 1 and 10 in culture, but these changes were not statistically significant (P = 0.654, n = 62; P = 0.313, n = 59, respectively; all the preceding reported statistical tests were performed using the Kruskal-Wallis one-way ANOVA on ranks). We did not measure the capacitance in most of the neurons in which gA and gh were measured. However, an increase of conductance density of these two currents with age in culture is not likely because the average capacitance during this period increased by the same amount as these conductances (25%). Surprisingly, the increase in capacitance during the first week in culture (cm on day 1 = 0.437 ± 0.154 nF, cm on day 7 = 0.548 ± 0.238 nF) was not statistically significant (P = 0.106, n = 87) similar to what was observed by Turrigiano et al. (1995). Furthermore, the electrotonic length of these neurons is relatively short (thus neurons are electrotonically compact) and not significantly different between day 1, when there is virtually no growth (L = 1.67 ± 0.61, n = 16), and day 6, when extensive growth is apparent (see Fig. 2, B–D; L = 1.73 ± 0.781, n = 8, P = 0.842, Student’s t-test). From these results, we conclude that neurons at these stages grow by extensively stretching the existing membrane rather than by incorporating significant amounts of new membrane.
To determine whether the conductances changes described in the preceding text correlate with STG neuron identity, we recorded gCa from seven pyloric and seven gastric neurons, ages 1 and 6 days in culture. We found no statistically significant difference between pyloric and gastric neurons (P = 0.841, Student’s t-test). However, we found a statistically significant difference between ages 1 (0.007 ± 0.002 µS) and 6 days (0.024 ± 0.014 µS; P = 0.012, Student’s t-test) of the pooled data, confirming the results described in the preceding text for nonidentified cells. We also measured gK, gA, and gh from 30 additional identified neurons (16 pyloric neurons and 14 gastric neurons). We grouped all neurons aged 5–7 days in culture and compared pyloric versus gastric neurons. No statistically significant difference in any of the three conductances between the two cell types was observed (P = 0.366, 2-way ANOVA), with gK = 0.30 ± 0.34 µS (pyloric) versus 0.23 ± 0.15 µS (gastric), gA = 0.93 ± 0.65 µS (pyloric) versus 0.77 ± 0.53 µS (gastric), gh = 0.020 ± 0.009 µS (pyloric) versus 0.018 ± 0.008 µS (gastric).
Activity changes induced by patterned stimulation
The changes in activity patterns and the accompanying conductance changes described in the preceding text, as well as previous observations in cultured lobster STG neurons (Turrigiano et al. 1994), suggest that isolated STG neurons follow a set course of spontaneous conductance changes and consequent modifications of activity that may be genetically predetermined. However, although a neuron may be on a predetermined course to ultimately become an oscillator, it may also be able to modify its pattern of activity as a function of the inputs it may receive (Cudmore and Turrigiano 2004; Franklin et al. 1992; Garcia et al. 1994; Golowasch et al. 1999a; Li et al. 1996; Turrigiano et al. 1994). We tested this possibility in our crab STG neurons by rhythmically stimulating them with current pulses and measuring possible changes in their patterns of activity. We found that in response to stimulation with hyperpolarizing current pulses (to bring the Vm from the resting potential to approximately –120 mV), the majority of cells (60%) did not change their activity pattern (Fig. 4D), remaining silent (28%), tonic (12%), or oscillatory (20%). We refer to this as no change in excitability. Similar to previous observations in lobster STG neurons (Turrigiano et al. 1994), we observed a small set of oscillatory neurons (10%) that reduced their excitability to tonic firing (Fig. 4, B and D). We were surprised to find that 30% of the stimulated neurons switched from either silent to oscillatory (26%, Fig. 4, A and D) or tonic to oscillatory activity (4%, Fig. 4, C and D) because this was not observed by Turrigiano et al. (1994) in their study of lobster STG neurons. We assume these changes in activity to reflect an enhancement of neuronal excitability. We observe no difference in the resting potential, Vrest, of neurons that change activity with stimulation (Vrest = –69.1 ± 8.9mV, n = 11) from those neurons that do not change activity (Vrest = –65.5 ± 3.8mV, n = 11, P = 0.397, unpaired Student’s t-test). Each set of neurons produced a slight but statistically significant depolarization in response to patterned stimulation (neurons with change: 
Vrest = +6.1 ± 5.8 mV, P < 0.001; neurons without change: Vrest = +6.6 ± 6.6 mV, P < 0.001, unpaired Student’s t-test). This depolarization, however, is indistinguishable between these two groups (P = 0.525, n = 22, unpaired Student’s t-test). Furthermore, we see no difference in the depolarization whether cells increased their excitability (silent or tonic to oscillatory transitions, Fig. 4D) or decreased their excitability (oscillatory to tonic transitions, Fig. 4D, P = 0.958, unpaired Student’s t-test). From this we conclude that the slight depolarization observed in stimulated cells is a nonspecific effect of stimulation.
In contrast to what was shown by Turrigiano et al. (1994), we found no consistent correlation of any of these activity changes with the presence of a measurable postinhibitory rebound (PIR) in these neurons. Figure 4E shows the stimulation-induced membrane potential changes during the "beginning" and "end" of the stimulation period used to induce activity changes. The traces marked A–C correspond to the cells the results of which are shown in A–C in this figure. The last trace, showing a relatively large PIR capped with an action potential, corresponds to an oscillatory neuron the activity of which did not change with stimulation. In fact, most of the stimulated crab STG neurons shown in Fig. 4 (59%) express no measurable PIR at all (see traces A and C in Fig. 4E). We found that neurons that do not show a change in activity induced by patterned stimulation generate a PIR on average almost twice as large (4.5 ± 7.2 mV, n = 30) as those that do (2.7 ± 3.5 mV, n = 16), but this difference is not statistically significant (P = 0.363, unpaired Student’s t-test). As can be seen in all four examples shown in Fig. 4E, there was no major change in PIR properties or amplitude during the course of the stimulation.
Interestingly, none of the recorded neurons showed a transition from a silent to a tonic (or from a tonic to a silent) pattern of activity, suggesting a lack of sensitivity to patterned stimulation of those currents responsible for the generation of action potentials. Alternatively, in these cells some of the intermediate signaling molecules leading from the detection of membrane potential changes to the modification of the ionic currents responsible for the changes in activity may be absent. In all cases where recordings could be held long enough (2–4 h), reversal of the activity pattern was always significantly slower than the induction phase and almost always partial. We rarely observed a complete reversal of these effects. Figure 4, B and C, shows the only two examples (of 28 cases) we obtained of almost complete reversal of activity, and in both it took 2.5 h to complete.
It is important to note that the ability for neurons to regulate activity in an activity-dependent manner did not appear to be related to the age of the neurons in culture but rather to their activity state at the time of stimulation. A similar fraction (40%) of "young" neurons (ages 1–4 days in culture) and "old" neurons (31%, ages 5–8 days in culture) could be induced to change their state of activity. However, most stimulated tonic (average age = 5.3 days) and oscillatory (average age = 3.5 days) neurons were older than stimulated silent (average age = 1.1 days) cells simply because of the spontaneous progression in activity observed in culture (see Fig. 3A).
Within the relatively simple activity pattern categories we have classified these neurons into, there is a wide range of variability in terms of action potential frequency, duration, and amplitude, slow-wave oscillation frequency and amplitude, threshold current required to elicit patterns of activity, etc. We reasoned that this variability may be related to the expression of large, variable amplitude outward currents (Golowasch et al. 1999a) and that reducing their amplitude may make the activity patterns more uniform and reveal a more consistent effect of rhythmic stimulation on neuronal activity. When outward K+ current amplitudes were reduced with 20 mM TEA in the bath, we did indeed observe a reduction in the variability of the activity states (Fig. 5). As shown before by Turrigiano and Marder (1993) in lobster isolated STG neurons, in the presence of TEA, most crab STG neurons showed a tendency to generate slow oscillations with relatively long depolarizations (Fig. 5B, control, n = 9/15) while all others were silent (Fig. 5A, control, n = 6/15) before rhythmic stimulation was begun. When these neurons were rhythmically stimulated, 100% of the initially silent neurons developed slow and large-amplitude oscillations (Fig. 5A, after stim), 100% of the initially oscillatory neurons increased the duration of the slow depolarizations by 83% from 319 ± 138 to 584 ± 369 ms (P = 0.030, n = 9, Fig. 5B, after stim), increased their amplitude by 40% from 24.4 ± 14.3 to 34.1 ± 14.2 mV (P < 0.001, n = 8, Fig. 9B, left), and also increased their oscillation period by 170% from 473 ± 314 to 1284 ± 936 ms (P = 0.046, n = 6, see Figs. 5B and 9B, left). Period was calculated on longer recordings than shown in Fig. 5. Comparisons were made with unpaired Student’s t-tests.
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) and (see Effect of stimulation protocol) the characteristics of the input. To determine if the different activity changes observed in response to patterned stimulation correlate with neuronal identity, we looked at the responses of identified pyloric versus gastric neurons in the presence of 20 mM TEA. Of a total of eight pyloric neurons and three gastric neurons, all expressed oscillatory activity. We measured oscillation amplitude (32.1 ± 19.2 mV), cycle period (590 ± 450 ms), and duty cycle (0.71 ± 0.28) and measured stimulus-induced changes in these three measures. We found no statistically significant difference in the changes induced by stimulation between pyloric and gastric cells in any of these three measures (P = 0.411, 2-way RM ANOVA).
Conductance changes induced by patterned stimulation
In neurons stimulated with rhythmic hyperpolarizing pulses, we observed a statistically significant decrease in high-threshold K+ conductance, gK, recorded at +10 mV in normal saline of 45% (0.58 ± 0.07 µS before stimulation, 0.31 ± 0.04 µS after stimulation, P = 0.005, n = 6, paired Student’s t-test; Fig. 4F, left). In those neurons in which the activity was not modified, no change in gK was observed (0.65 ± 0.17 µS before stimulation; 0.68 ± 0.15 µS after stimulation, P = 0.375, n = 6, paired Student’s t-test; Fig. 4F, right). These results are consistent with an increased excitability of those neurons sensitive to prolonged stimulation as would be expected for neurons that switched activity from a silent or tonic pattern to an oscillatory pattern (Fig. 4, A and C). These results appear to be inconsistent with the change in activity from oscillatory to tonic firing that we saw in a small subset of stimulated neurons (12%, Fig. 4D), which under our definition, would be considered due to a reduction of excitability (see dynamic clamp results below and DISCUSSION).
STG neurons in situ (cf. Golowasch and Marder 1992; Graubard and Hartline 1991) and in culture (Turrigiano et al. 1995) express large outward currents dominated by a TEA-sensitive IK(Ca). In contrast with the complete block of IK(Ca) by TEA in situ (Golowasch and Marder 1992; Graubard and Hartline 1991), TEA has been reported to incompletely block IK(Ca) in cultured STG neurons (Hurley and Graubard 1998). In our hands, 20 mM TEA eliminates 84 ± 6% (n = 6) of the total high-threshold outward current in cultured STG neurons. IK(Ca) constitutes 20% of the outward current recorded in 20 mM TEA (defined as the current additionally blocked by 200 µM Cd2+; the high-threshold current remaining in the presence of Cd2+ corresponds to a delayed rectifier current, IKd). In the presence of 20 mM TEA, we observed that prolonged patterned stimulation induced a significant reduction of the outward current for all voltages above –20 mV (P = 0.004, n = 6, 2-way RM ANOVA, Fig. 6A) but with no apparent effect on the voltage dependence of activation (Fig. 6A; V1/2 before stimulation = –9.4 ± 17.6 mV, V1/2 after stimulation = –10.4 ± 14.7 mV, P = 0.900, n = 6, paired Student’s t-test). We estimated the maximum conductance from the current averages measured at 0 mV. Before stimulation, the estimated maximum conductance was 0.12 ± 0.06 µS, which decreased by a statistically significant 22% to 0.09 ± 0.06 µS (P = 0.022, n = 7, paired Student’s t-test). In the absence of known specific K+ current inhibitors, these data suggest that most of the spontaneous and stimulation-induced changes of IK could be attributed to IK(Ca) because 87% of the total IK corresponds to IK(Ca), as defined by K+ current blockade with Cd2+ (Golowasch and Marder 1992), and 13% to a delayed rectifier K+ current. Thus the 45% stimulation-induced reduction in the total K+ current that we observe cannot be accounted for by effects on the delayed rectifier only. Furthermore, because we observe a stimulation-induced reduction of 22% of the remaining total outward current in the presence of 20 mM TEA and 20% of that current can be further blocked with Cd2+ (and thus corresponds to IK(Ca)), we conclude that an effect of rhythmic stimulation exclusively on IK(Ca) is in principle sufficient to account for all our observations of activity-dependent effects on outward currents.
We performed similar measurements on six identified pyloric neurons and five identified gastric neurons to confirm if any of these changes are neuron type-specific. We compared the high-threshold K+ current I-V relationship difference (after normalization) before and after 0.33-Hz hyperpolarizing stimulation for pyloric and gastric neurons separately as was done with unidentified neurons before (see Fig. 6A, bottom). A two-way RM ANOVA reveals no statistical significant difference in the effect of rhythmic stimulation between identified pyloric and gastric neurons (P = 0.546).
We isolated ICa as described in METHODS (Fig. 6B). Hyperpolarizing stimulation for as little as 15 min induced a marked increase in ICa (Fig. 6B, top). Figure 6B also shows the peak current-voltage relationship before (control, 
) and after rhythmic stimulation of STG neurons (after stim, 
). A statistically significant increase for all voltages above –20 mV is observed (P = 0.011, n = 15, 2-way RM ANOVA followed by Bonferroni post hoc analysis), with the peak conductance measured at +10 mV growing 2.5-fold from 0.014 ± 0.031 to 0.036 ± 0.031 µS. (P = 0.046, n = 7, paired Student’s t-test). We observed no significant effect of stimulation on the voltage dependence of activation of this current (Fig. 6B; the half-maximal activation voltage, V1/2 before stimulation = –2.5 ± 10.0 mV, V1/2 after stimulation = –5.5 ± 30.2 mV, P = 0.099, n = 7, paired Student’s t-test).
As can be seen in the current traces in Fig. 6B (top), ICa measurements are slightly contaminated with IK, although the very early measurements of ICa minimized this contamination. In 20 mM TEA 20% of that K+ current corresponds to IK(Ca), the conductance of which we argue is reduced by rhythmic hyperpolarizing stimulation. To eliminate this possibility, we first tested the effect of 100 mM TEA on IK. We found that the cells were not adversely affected by this extremely high concentration. We also found that adding 200 µM Cd2+ does not produce an additional block of IK (not shown). We take this as evidence that 100 mM TEA completely eliminates IK(Ca), leaving intact only part of the delayed rectifier conductance. As a further way to isolate ICa, we measured the Ca2+ tail currents (in the presence of 100 mM TEA + 0.1 µM TTX) at –80 mV after activating ICa with 800-ms-long depolarizing pulses in the range –60 to +30 mV (Fig. 6C). The K+ equilibrium potential in these neurons is close to –80 mV. Thus these tail currents should be almost entirely free of contaminating K+ currents. Under these conditions, we observe a highly significant increase in ICa at all voltages above –20 mV (P = 0.001, n = 5, 2-way RM ANOVA followed by Bonferroni post hoc analysis). At +20 mV, we calculate a highly significant Ca2+ conductance increase from 0.040 ± 0.047 to 0.061 ± 0.056 µS (P < 0.001, n = 10, paired Student’s t-test).
In contrast with the effect on IK and ICa, the leak conductance showed no change in conductance in these experiments (0.004 ± 0.003 µS before stimulation and 0.005 ± 0.002 µS after stimulation, P = 0.712, n = 7, paired Student’s t-test). Leak conductance was determined in these experiments from the current changes elicited in response to the voltages steps from –40 to –50 (or –60) mV. Additionally, the transient IA current (measured in TEA to minimize other K+ currents and with IK and leak current subtracted) is completely unaffected by patterned stimulation in cultured crab STG neurons (Fig. 6D, P = 0.918, 2-way RM ANOVA over the activation range: –40 to +30 mV). Specifically, the average peak conductance gA, V1/2, and s values measured at +10 mV were gA = 0.32 ± 0.11 µS, V1/2 = –24.9 ± 5.6 mV, and s = 9.5 ± 3.5 mV before stimulation, and gA = 0.33 ± 0.13 µS, V1/2 = –23.5 ± 5.3 mV, and s = 7.9 ± 2.4 mV after stimulation were likewise not statistically significantly affected by stimulation (P = 0.938, n = 7, 2-way RM ANOVA).
The hyperpolarization-activated current’s voltage-dependence (i.e., steady-state activation curve) is strongly shifted to more negative values than those observed in cultured lobster STG neurons or crab’s LP neuron in situ (Golowasch 1992 #11; Turrigiano et al. 1995) before (V1/2 = –104.1 ± 4.4 mV, slope factor s = 8.3 ± 3.7 mV, n = 7), and neither of these values nor the maximum conductance were affected by patterned stimulation. The maximum conductance measured at –120 mV was 0.015 ± 0.007 µS before stimulation, and 0.018 ± 0.010 µS after stimulation (P = 0.140, n = 8, paired Student’s t-test).
Role of calcium influx in activity-dependent regulation of conductances
The conductance changes reported in the preceding text most likely occurred in response to the experimentally imposed activity pattern and were not due to effects of the electrode-filling solutions (K-citrate, K2SO4, or TEA.Cl + CsCl) or of different bathing solutions (TEA, TTX, Cs+), and occurred in the absence of any known growth factors or neuromodulators. For activity to be responsible for these changes, neurons need to be able to detect changes in their own patterns of activity. A plausible candidate for such a gauge of activity is intracellular Ca2+ (Bito et al. 1997; De Koninck and Schulman 1998; Liu et al. 1998; Schulman et al. 1995). Indeed, in our neurons the only conditions that block the effects of rhythmic stimulation are those that interfere with Ca2+ influx (neither the K+ current blocker TEA nor the Na+ current blocker TTX do). Figure 7A shows peak ICa recorded in 100 mM TEA and C shows IK recorded in 20 mM TEA, before (control) and after 0.33-Hz hyperpolarizing stimulation in the presence of 200 µM Cd2+ (stim in Cd2+). Cd2+ is known to block the high-threshold ICa in STG neurons (Golowasch and Marder 1992; Graubard and Hartline 1991; Turrigiano et al. 1995), and we reasoned that stimulation in the presence of a blocker of Ca2+ influx should eliminate the effect of stimulation on IK and ICa. Indeed we observed that 0.33-Hz hyperpolarizing stimulation in the presence of Cd2+ eliminates the enhancing effect on ICa (P = 0.308, n = 5) and the depressing effect on IK (P = 0.618, n = 5, 2-way RM ANOVA). The inset in Fig. 7A (re-stim in Ca2+) also shows a strong enhancement of ICa recorded in one (of 2) cells in which we succeeded to wash out Cd2+ and further stimulate with hyperpolarizing pulses for 30 additional minutes with Ca2+ influx restored. Furthermore, Fig. 7B shows an inward current that shows virtually no inactivation when Ca2+ is replaced with Ba2+ in the extracellular solution. With Ca2+ influx thus minimized, no change in the amplitude of the inward current now carried by Ba2+ (control) was observed in response to prolonged patterned stimulation (after stim, Fig. 7B). No significant changes were recorded over the voltage-dependent activation range (–30 to +30 mV) of this current (P = 0.300, n = 5, 2-way RM ANOVA).
Effect of stimulation protocol
The results shown thus far suggest that rhythmic neuronal activity plays an important role in determining both activity and ionic conductance changes. The stimulation protocol used thus far has been hyperpolarizing 1-s-long pulses every 3 s (METHODS). Does the effect of rhythmic stimulation on activity and ionic currents depend on the properties of the stimulus? We tested this by measuring the high-threshold K+ current, IK (Fig. 8). Cells were hyperpolarized rhythmically at a slower rate (8 s pulses every 9 s, i.e., 0.11-Hz Hype, Fig. 8A) and at a slightly faster rate (1-s pulses every 2 s, i.e., 0.5 Hz, Fig. 8B) than the standard stimulation frequency of 0.33 Hz. Because our several controls on identified pyloric and gastric neurons have shown no correlation of any of the measurements of activity or conductance with neuronal cell type, we performed this experiment on unidentified neurons. Figure 8A shows the effects on IK of applied 0.11-Hz hyperpolarizing stimulation for 25 min, followed by 35–45 min of standard 0.33-Hz hyperpolarizing stimulation. Compared with the results shown in Fig. 6A, where 0.33-Hz stimulation induces a clear and statistically significant decrease in amplitude of IK at all voltages above –40 mV, 0.11-Hz hyperpolarizing stimulation instead induces a statistically significant increase of the current amplitude at voltages 
0 mV (P = 0.002, n = 6, 2-way RM ANOVA; Fig. 8A, solid gray triangles). When 0.11-Hz hyperpolarizing stimulation was followed by 0.33-Hz stimulation (Fig. 8A, open gray squares), we observed a statistically significant decrease of the current amplitude at voltages above –20 mV (control vs. 0.33 Hz stim: P = 0.011; 0.11-Hz Hype vs. 0.33-Hz hype: P < 0.001, n = 6, 2-way RM ANOVA) comparable to our previous observations (Fig. 6A). To establish if this is a specific effect on the high-threshold K+ current, we measured the effects of the same stimulation sequence on IA, which we have already shown to be unresponsive to 0.33-Hz stimulation (Fig. 6D). No statistical significant effect of either stimulus pattern was observed on IA (P = 0.869, n = 5, 2-way RM ANOVA). On the other hand, only a slightly higher hyperpolarizing stimulation frequency (0.5 Hz) compared with the standard 0.33-Hz frequency results in no significant change in amplitude of IK at any voltage (Fig. 8B, P = 0.779, n = 5). IA was also not affected by 0.5-Hz hyperpolarizing stimulation (P = 0.254, n = 5, 2-way RM ANOVA). The fact that hyperpolarizing stimulation within a voltage range in which no Ca2+ current has been directly detected in STG neurons in culture or in situ raises the question of what the path may be for cytoplasmic Ca2+ increase. Because ICa is strong at –10 mV, we stimulated STG neurons with depolarizing pulses from –40 to –10 mV in voltage clamp to ensure Ca2+ influx through a path known to be present in these cells. Figure 8C shows the results of such an experiment. Depolarizing stimulation had no discernible effect on IK (P = 0.526, n = 5, 2-way RM ANOVA). To demonstrate that this result is specific to depolarizing stimulation, in two instances, we were able to maintain the cells impaled long enough to stimulate them with 0.33-Hz hyperpolarizing pulses for additional 40 min. Figure 8C, inset, shows that the characteristic downregulating effect of hyperpolarizing stimulation could be clearly elicited in this neuron. Finally, depolarizing stimulation also had no significant effect on IA (P = 0.898, n = 5, 2-way RM ANOVA).
Additionally, as would be expected from the effects of 0.11-Hz hyperpolarizing stimulation on IK mentioned in the preceding text, 0.11-Hz stimulation had a statistically significant effect on the amplitude of the slow oscillations recorded in the six neurons that we stimulated (50.2 ± 14.0 mV before stimulation, 31.7 ± 17.8 mV after stimulation; P = 0.016, Student’s t-test). We could not measure either burst duration or cycle period because 0.11-Hz stimulation had the additional effect of drastically reducing the number of oscillations measured with a current pulse of any levels, in most cases to only one or zero oscillations. These results, although not quantifiable, are consistent with an enhancement of a high-threshold K+ current.
Dynamic-clamp experiments
The effects of prolonged stimulation on voltage-dependent currents and neuronal activity only partially reversed during the time we could maintain the recordings (2–4 h). To verify if the spontaneous and stimulation-induced conductance changes (IK decrease and ICa increase) are indeed sufficient to account for the observed alterations in neuronal patterns of activity, we introduced negative or positive outward and inward conductances with dynamic clamp. We fitted with Hodgkin and Huxley-type equations the high-threshold K+ current difference and separately the Ca2+ current difference of neurons sensitive to patterned stimulation in normal saline. For these fits, we chose neurons that responded to patterned stimulation with high-threshold K+ and Ca2+ current changes close to the population average. The best fit to the K+ current could be obtained with two conductance components, one transient (gKtr) and one sustained (gKst), whereas the Ca2+ current could be fitted well with a single component (gCa). The conductance parameters thus obtained and used for our dynamic-clamp experiments are given in Table 1. Because the voltage dependence of these currents does not appear to be affected by patterned stimulation, we modified only the maximum conductances to mimic the changes observed on these two currents. Figure 9A shows the results of one such experiment on naïve unstimulated neurons recorded in normal saline. Activity was elicited by small depolarizing current injections. Conductance values indicated above the arrows correspond to the maximum conductance values of each dynamic-clamp current. Many different patterns of activity could be produced by relatively small changes of the maximum conductance values, which depended on the particular neuron recorded (Fig. 9A). However, we could repeatedly induce oscillatory activity by reducing the outward currents (negative conductance) and/or increasing the inward current (positive conductance, Fig. 9A, top and middle). It was relatively easy to also find combinations of maximum conductance values within the range of the spontaneous or stimulation induced conductance changes described before that could produce tonic firing in a neuron that was originally oscillatory (Fig. 9A, bottom). Figure 9B shows in more detail that some properties induced by prolonged rhythmic stimulation cannot only be mimicked by adding gCa and subtracting gK but also reversed by subtracting gCa and adding gK after they were induced by stimulation. Before rhythmic stimulation was started, the neuron shown in Fig. 9B (top left) was bathed in 20 mM TEA to induce slow, large-amplitude oscillations. Adding gCa and subtracting gK with dynamic clamp increased the amplitude and slightly decreased the frequency of oscillations (Fig. 9B, top right). A similar but more pronounced effect was later observed after rhythmically stimulating the neuron for 20 min with dynamic clamp discontinued. Finally, the enhanced oscillation amplitude and reduced oscillation frequency induced by rhythmic stimulation was partially reversed by subtracting gCa and adding gK with identical levels as used immediately before to induce the changes with dynamic clamp (Fig. 9B, bottom right).
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; Desai et al. 1999; Franklin et al. 1992; Galante et al. 2001; Hong and Lnenicka 1995; Li et al. 1996; Linsdell and Moody 1994; Luther et al. 2003; Turrigiano and Nelson 2004; Turrigiano et al. 1994). Here we have used adult cultured STG neurons of the crab C. borealis to identify mechanisms that underlie the recovery and stabilization of rhythmic activity after dissociation. We find that downregulation of the high-threshold K+ current, IK, and upregulation of a Ca2+ current, ICa, is called into play during the spontaneous regulation of activity after dissociation. Prolonged 0.33-Hz rhythmic stimulation with hyperpolarizing pulses induces similar changes in the same two currents. However, low-frequency hyperpolarization (0.11 Hz) had the opposite effect on IK and slightly higher frequency (0.5 Hz) had no effect at all (Fig. 8A), indicating that crab STG neurons are sensitive to the temporal properties of the stimulus as previously suggested by Turrigiano and collaborators. As a consequence STG neurons may be tuned to the oscillating frequencies of the network each of these neurons are normally part of (Turrigiano et al. 1994). From this it would be expected that neurons belonging to networks the natural frequency of which is not the same would respond differently to the same pattern of stimulation or, alternatively, develop a different frequency of oscillation in culture. Surprisingly, however, we found that identified neurons belonging to both the pyloric and the gastric mill networks, which typically oscillate in situ at 5- to 10-fold different frequencies, developed oscillations in culture with indistinguishable frequencies. We further found that gastric and pyloric neurons cannot be distinguished during the culture period studied (10 days) on the basis of their activity changes or the ionic mechanism involved in their responses to stimulation. Preliminary ongoing experiments indicate that in culture pyloric neurons appear to retain the properties developed during the first week in culture, whereas gastric neurons appear to slowly differentiate from a pyloric neuron phenotype in a time scale of weeks via an unknown mechanism.
We find that excitability cannot only be reduced by patterned stimulation as has previously been reported (Turrigiano et al. 1994) but can also be heightened with the same type of stimulation, an excitability increase being three times more likely in crab neurons than an excitability reduction. The apparent difference in response to rhythmic stimulation of lobster neurons, which in the study of Turrigiano and collaborators were not shown to be able to enhance their excitability, may stem from the fact that in their study only bursting neurons were selected for stimulation and analysis (Turrigiano et al. 1994). We find that oscillating neurons can only respond to stimulation by switching to either tonic firing (similar to lobster neuron responses) or not at all. In their study, Turrigiano et al. (1994) apparently did not analyze bursting neurons that did not respond to stimulation.
Spontaneous activity changes
The progressive decrease in the proportion of silent STG neurons in primary culture and the increase in the proportion of neurons expressing tonic and, later, mostly oscillatory activity, suggests a predetermined tendency to oscillate and the existence of homeostatic mechanisms that allow neurons to restore their oscillatory activity after it is lost (see also Turrigiano et al. 1995). Pyloric network neurons can behave as oscillators when acutely isolated from the network but, with the exception of the single pacemaker AB neuron, all oscillate irregularly (Bal et al. 1988). These oscillations occur only when neuromodulatory inputs to the STG are intact (Nusbaum and Beenhakker 2002). In the absence of these neuromodulatory inputs, all STG neurons, including the AB neuron, lose the ability to oscillate (Bal et al. 1988). Our results and those of Turrigiano et al. (1995) indicate that most (if not all) STG neurons in time evolve robust oscillatory activity when isolated from network and from neuromodulatory influences.
In our culture conditions, neuromodulators are completely absent and the recovery of rhythmic activity of isolated STG neurons is reminiscent of the recovery of rhythmic activity in the intact pyloric network after the permanent removal of neuromodulatory inputs (Luther et al. 2003; Thoby-Brisson and Simmers 1998). In lobsters, this latter recovery is accompanied by the development of oscillatory properties by some identified pyloric network neurons within a few days (Thoby-Brisson and Simmers 2002). Thus the acquisition of oscillatory properties we and Turrigiano et al. (1995) have observed in isolated STG neurons in culture provides a plausible mechanism for the recovery of rhythmic activity in the network. It is interesting, however, that the conductances that have been shown to be spontaneously regulated in the intact network after decentralization are not the same as those regulated in dissociated neurons in culture (Thoby-Brisson and Simmers 2002) (see Ionic mechanism of activity regulation).
Two hypotheses (as of yet not tested) may explain the suppression of endogenous oscillatory properties in neurons within the intact STG. One, oscillatory properties may be suppressed by neuromodulators exerting trophic effects (Le Feuvre et al. 1999; Thoby-Brisson and Simmers 2000). Two, activity itself may regulate the expression of endogenous oscillatory properties (Golowasch et al. 1999b; Turrigiano et al. 1994). Our results are consistent with both hypotheses. In the intact network, neuromodulators may somehow suppress the expression of endogenous oscillatory properties in most neurons. Once this input is eliminated (by cell dissociation or by removal of neuromodulatory inputs in situ), constraints on the expression of these properties are removed and oscillatory activity can be expressed by up- and downregulation of ionic currents such as those characterized in this work and those characterized previously by Turrigiano et al. (1995). Spontaneous emergence of oscillatory properties in culture conditions may be genetically determined as suggested by the spontaneous changes in activity and ionic currents described in this study and by Turrigiano et al. (1995). However, this possibility does not exclude the possibility that activity itself may also regulate neuronal electrical properties (Desai et al. 1999; Golowasch et al. 1999a; Liu et al. 1998).
Stimulation-dependent effects on activity
Our results show that isolated STG neurons are sensitive to prolonged rhythmic stimulation. Neurons displaying either silent or tonic firing activity can be induced to oscillate, whereas neurons that show oscillatory activity remain able to revert to tonic firing (but not to the silent) states (Fig. 4).
The large variability of the effects on activity of prolonged stimulation (Fig. 4D) may partly be due to the heterogeneity of our sample of (mostly unidentified) neurons. However, when outward currents were reduced in the presence of TEA, 100% of the stimulated neurons increased their excitability (Fig. 5). Furthermore, 100% of the neurons in which outward currents were measured in the presence of TEA showed a decrease of the remaining TEA-resistant IK, and 100% of those neurons in which ICa was measured showed an increase in conductance. The homogeneity of these effects, although variable in their extent, suggests to us that all STG neurons, irrespective of cell type, can regulate their excitability in an activity-dependent manner and via the same ionic mechanism, namely by downregulation of a IK current and an upregulation of ICa. This conclusion is further supported by the similarity of the results of our measurements of change in IK in identified pyloric and gastric neurons.
Ionic mechanism of activity regulation
Only two ionic currents appear sensitive to prolonged patterned stimulation in dissociated C. borealis STG neurons, ICa and IK. The same two currents are responsible for the spontaneous changes of activity seen in these neurons. It has previously been shown in cultured lobster STG neurons that, in addition to these currents, changes in two Na+ currents also correlate with spontaneous changes of activity from silent to tonic firing but that only ICa continues to increase as neurons become bursters (Turrigiano et al. 1995). We cannot exclude the participation of Na+ currents in cultured crab STG neurons. However, tonic firing becomes gradually less and less prevalent among neurons recorded up to day 6 in culture (Fig. 3A) and almost completely absent after 3 wk in culture (not shown). Also, silent cells could never be induced to fire action potentials with prolonged hyperpolarizing (patterned or constant) stimulation. These observations suggests that Na+ currents in crab STG neurons, apparently in contrast with their lobster counterparts, may not be subject to long-term upregulation or to activity-dependent regulation but may only spontaneously and transiently be upregulated during the first few days in culture. Additionally, Turrigiano et al. (1995) observed a drastic reduction in the transient A current, IA, as neurons transition between silent, tonic, and oscillatory states. In contrast, we have seen no change in IA at any time in culture (Fig. 3C), and also no changes in IA are induced by prolonged stimulation (Fig. 6D and see RESULTS). The lack of effects of prolonged hyperpolarizing stimulation on IA in crab cultured STG neurons is consistent with the lack of effects of similar hyperpolarizing stimulation on IA in two different identified crab pyloric neurons in situ (Golowasch et al. 1999a). However, although IA was strongly affected by depolarizing stimulation of IC neurons in situ (Golowasch et al. 1999a), IA was not affected in any neurons in culture (this study). On the other hand, in situ hyperpolarizing stimulation of these same cells did also not affect the high-threshold K+ currents (Golowasch et al. 1999a) that we find to be affected in our STG cultured neurons. Finally, the hyperpolarization-activated current Ih did not change either spontaneously or in response to stimulation in our study, but it does increase spontaneously after decentralization in lobster PD neurons [(Thoby-Brisson and Simmers 2002); it was not measured in lobster cultured neurons (Turrigiano et al. 1995)]. It could be argued that these differences in spontaneous changes and response to rhythmic stimulation may stem from the fact that our cultured neurons were not precisely identified to the level of individual cell types (but only to the level of network identity). Although this is possible, we find that, as stated in the previous section, all the neurons stimulated respond, and all respond in the same manner at the level of ionic conductance changes, suggesting that neuronal identity appears not to be a factor. It is possible that in situ the high-threshold K+ currents sensitive to stimulation represents a small current relative to the total high-threshold K+ current and thus activity changes, or total conductance changes, could not be unambiguously discerned. Alternatively, the process of dissociation in which cells lose most of their dendritic structure, the special culture conditions (artificial saline, complete absence of neuromodulators, etc) and the absence of synaptic network interactions may reveal a new, or alter an existing activity-sensing mechanism.
The 2.5-fold stimulation-induced ICa increase was achieved with 30–60 min of stimulation (Fig. 6B). ICa increases spontaneously also by 2.5-fold but over 4 days in culture (Fig. 3B). Similarly, the high-threshold K+ conductance was reduced by 45% in normal saline with 45–60 min of rhythmic stimulation but changes spontaneously by only 25% over 6 days in culture. Thus although the same currents appear targeted during spontaneous and induced activity changes, these effects may occur via two different but probably overlapping pathways. Slow, spontaneous changes may predominantly require gene transcription and protein synthesis, similar to the transcription requirement for spontaneous but slow pyloric activity recovery in the intact pyloric network after the permanent removal of neuromodulatory inputs (Thoby-Brisson and Simmers 2000).
Our dynamic-clamp experiments confirm that an increase in Ca2+ conductance, gCa, and a decrease in K+ conductance, gK, are sufficient to explain all the observed changes in activity. We were able to induce a switch from either silent or tonic firing to oscillatory activity by modifying the same conductances (gCa, gK), within the conductance ranges observed physiologically, that we showed to be affected during spontaneous activity changes and by patterned stimulation (Fig. 9A). Although relatively small gK changes were sufficient to generate significant activity changes, the gCa levels we needed to use to induce activity changes were comparable to the gCa changes observed in response to prolonged patterned stimulation. Although increasing gCa with dynamic clamp always increased the tendency of neurons to burst, it is important to remember that this current is not a Ca2+ current in the biological sense because no Ca2+ influx occurs. An increase in Ca2+ influx due to an augmented biological gCa will be accompanied by an increase in IK(Ca), which can reduce the excitability of a neuron and thus induce tonic firing. However, even a dynamic-clamp-reduced K+ current in a background of high ICa can switch an oscillatory to a tonically firing neuron (Fig. 9A, bottom), similar to 41% of our rhythmically stimulated oscillatory neurons (Fig. 4, C and D). Consistent with our observations of stimulation-induced activity changes, we could never induce tonic firing in silent cells by manipulating gCa and gK. Unlike Thoby-Brisson and Simmers (2002), we observed no spontaneous or stimulation-induced changes in Ih and did not test the effects of this current with dynamic clamp. This current is well known to be involved in the generation of oscillatory activity (Forti et al. 2006; Maccaferri and McBain 1996; McCormick and Bal 1997; Noble et al. 1992) and may be responsible for the enhancement of oscillatory properties observed in decentralized lobster preparations (Thoby-Brisson and Simmers 2002). Our dynamic-clamp results show that changes in ICa and IK are sufficient to explain our observations but certainly do not exclude the possible participation of Ih under other conditions or in other species.
Our observations suggest that spontaneous changes in Ca2+ and high-threshold K+ currents likely play a central role in homeostatic recovery of rhythmic function in the decentralized pyloric network of the crab STG. Decreases in K+ currents as a result of decreased neuronal drive are common to most if not all systems in which a homeostatic recovery of excitability can be identified (Zhang and Linden 2003). Although among inward currents sometimes Na+ currents are also affected (Desai et al. 1999; Mee et al. 2004), more commonly Ca2+ currents are strengthened due to decreased activity (Chung et al. 1993; Garcia et al. 1994; Su et al. 2002) or reduced due to enhanced neuronal activity (DeLorme et al. 1988; Franklin et al. 1992; Hong and Lnenicka 1995; Li et al. 1996). The effects we have observed of very slow frequency hyperpolarizing stimulation (0.11 Hz) do not appear to be consistent with the concept of homeostatic regulation of function. Rather a different mechanism that regulates these same conductances in a frequency-dependent manner and with a faster time course than the spontaneous recovery appears to be active in parallel. A solid conclusion on this will need to await further experiments to rule out possible unphysiological effects of the strong hyperpolarization used during stimulation in our study.
Intracellular signaling
We show that activity-dependent Ca2+ and K+ conductance changes depend on Ca2+ influx into these cells. We suggest that a Ca2+-dependent intracellular mechanism modifies the conductances of both IK and ICa. Ca2+ influx appears to be crucial also in other systems in mediating activity-dependent conductance and activity changes during development (Spitzer et al. 2002) and in the adult (Zhang and Linden 2003). Several possible Ca2+-dependent mechanisms may be involved, including transcription regulation (Spitzer et al. 2002; West et al. 2002), ion channel downregulation (Klein et al. 2003), and posttranslational modifications (Cudmore and Turrigiano 2004), which can be sensitive to the exact pattern and path of Ca2+ entry (Bito et al. 1997; De Koninck and Schulman 1998; Dolmetsch et al. 1998; Fields 1994; Li et al. 1996). We observed no correlation of stimulation-induced activity changes with expression of post inhibitory rebound (PIR) properties. This lack of correlation of activity or conductance changes with the measured PIR may lie in that PIR is a rather indirect measure of Ca2+ influx. If contaminated with a simultaneous activation of IK, a small Ca2+ influx may still take place with little or no measurable PIR. Second, we observed the opposite effect on the levels of IK and on activity when the 0.33-Hz rhythmic pattern of hyperpolarizing stimulation was replaced with low-frequency (0.11 Hz) hyperpolarization, i.e., an increase instead of a decrease in IK (Fig. 8A) and a reduction in the amplitude of the membrane potential oscillations. This is consistent with the possibility that a PIR-like mechanism during which a small but sufficient influx of Ca2+ takes place as suggested by Turrigiano et al. (1994). Prolonged hyperpolarization would simply close Ca2+ channels that are open near the resting potential and maintain them closed longer, thus preventing Ca2+ influx into the cells and bringing intracellular Ca2+ levels below the levels of the unstimulated neurons. Although this mechanism is plausible, it is harder to reconcile with our observation that only a slightly higher frequency of hyperpolarizing stimulation completely eliminates the effect of patterned stimulation on IK (Fig. 8B). It is conceivable, however, that the kinetics of Ca2+ influx can sensitively interact with the kinetics of intracellular Ca2+ uptake and of intracellular activation of Ca2+-dependent molecules involved in the activity-sensing mechanism to yield a highly frequency-specific transduction mechanism (Dolmetsch et al. 1998; Gorbunova and Spitzer 2002; Soderling et al. 2001). A more serious challenge to this hypothesis comes from the results showing a complete lack of effect of depolarizing stimuli (that ensure Ca2+ influx through ICa) on IK (Fig. 8C). These results strongly suggest that it is not merely Ca2+ influx that determines the effect on these currents. An interaction of the localization of the voltage-sensitive plasma membrane channels required for Ca2+ influx and the intracellular localization of the appropriate transducing signaling molecules would be required. Such an interaction has been observed in many other systems (Hobai and Levi 1999; Parekh 2006; Wu et al. 1999). The question still remains: what is the path of Ca2+ entry? Some forms of L-type Ca2+ channel, although they typically carry a high-threshold current, have a finite conductance at membrane potentials low enough (i.e., –70 mV) to contribute to the resting potential (Smith et al. 1993). As Fig. 6C shows, however, we observe no inward current at voltages near or below the resting potential of cultured STG neurons (i.e., –60 to –40 mV). No direct report to date exists of Ca2+ current mediating a Ca2+ influx at low membrane potentials in STG neurons, but such an influx has been inferred from the volt-age dependence of synaptic release in these cells and from Ca2+ imaging data showing a Cd2+-sensitive low-threshold Ca2+ influx (Kloppenburg et al. 2001). A very-low-level Ca2+ influx via these Cd2+-sensitive channels could go undetected and could trigger Ca-induced Ca release from intracellular stores thus amplifying the Ca2+ signal. This mechanism would account for the sensitivity to Cd2+ at the same time as the absence of measurable transmembrane Ca2+ current but would require postulating two different Ca2+ channels, with distinct voltage dependence and cellular localization and both with similar Cd2+ sensitivity. With the exception of Cd2+ sensitivity, such distinct voltage dependence and cellular localization is not uncommon (cf. Ivanov and Calabrese 2006a,b). It has been shown that hyperpolarization-activated channels can be permeable to Ca2+ (Yu et al. 2004). Because most cultured STG neurons express an Ih this is a possibility that remains to be tested. Another candidate pathway for Ca2+ influx are gap junction-forming hemi-channels. Gap junctions are known to be permeable to Ca2+ (Saez et al. 1989), and many STG neurons form gap junctions. Hemi-channels may also be permeable to Ca2+ although to our knowledge this has not been established for innexins.
In conclusion, we suggest that, consistent with our observations and those of Turrigiano et al. (1995), adult STG neurons have a natural predetermined tendency to oscillate independent of neuronal type. Activity can, however, be regulated in an activity-dependent manner over this natural tendency. Only two ionic currents, ICa and IK, appear to be involved in both the spontaneous development of oscillations and in stimulation-induced changes of activity in crab STG neurons. Our dynamic-clamp experiments show that regulation of only these two currents is sufficient to produce different changes in activity. The spontaneous changes in ICa and IK can explain the homeostatic recovery of pyloric network activity after removal of neuromodulatory input to the ganglion in situ, whereas the rhythmic stimulus-induced conductance changes appear to be to some extent at odds with the concept of homeostatic recovery of function. We also conclude that, at least during approximately the first week in culture, no discernible differences in the process of recovery of activity or the mechanisms involved can be made between neurons belonging to the two main functional neural networks (pyloric and gastric) found in the STG. Differences in the generation of oscillatory properties in pyloric versus gastric neurons in situ may thus be due to either synaptic interactions within the STG or to intrinsic mechanisms operating at longer time scales than 1 wk after dissociation.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: J. Golowasch, Dept. Mathematical Sciences, NJIT, University Heights, Newark, NJ 07102 (E-mail: Jorge.P.Golowasch{at}njit.edu)
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ABSTRACT
INTRODUCTION
, –55 mV (left) and –63 mV (right).
, pre) and after 45–60 min of stimulation (
) and shown as above. P < 0.004, n = 6. IK was normalized by currents measured in control conditions at 0 mV. B, top: typical recordings of a Ca2+ current at 0 mV in Cancer saline containing 20 mM TEA, 0.1 µM TTX. The neurons were also TEA and Cs+ loaded to reduce K+ currents to a minimum. The transient inward current (control) increased in amplitude after 30 min of rhythmic stimulation (after stim). Middle: raw voltage trace used to elicit the control current trace above. 
0.33-Hz Hype, open gray squares). The pairwise statistical comparisons between the 3 treatments is shown as P values (n = 6). Pairwise statistical comparisons of the origin of the differences among the treatments used the Bonferroni post hoc test and is indicated as **P < 0.001; *P < 0.05. B: control current measurements were obtained before stimulation (control, black squares) and after stimulation with hyperpolarizing 1-s-long pulses at 0.5 Hz (0.5-Hz Stim, open gray squares). P = 0.779, n = 5. C: control current measurements were obtained before stimulation (control, black squares) and after stimulation in voltage-clamp with depolarizing 1-s pulses from –40 to –10 mV at the standard 0.33-Hz frequency (depol stim, open gray squares). P = 0.526, n = 5. In 2 occasions, depolarizing stimulation was followed by 40-min stimulation with the standard 1-s hyperpolarizing pulses at 0.33 Hz. Measurements of 1 of these cells is shown in the inset (then 
) trace was recorded after washout of Cd2+ and additional 40 min stimulation in normal Ca2+ conditions. Bottom: average I-V plots of the normalized peak ICa measured 25 ms after the onset of the depolarizing test pulse, recorded before stimulation (control) and after stimulation in (and after washout of) Cd2+ (Stim in Cd2+). P = 0.308, n = 5. Inset: example of a single cell recorded under the 3 conditions indicated above. B, top and middle: noninactivating IBa (top) elicited by a test pulse from –40 mV to +10 mV (middle) before (control) and after 45 min of rhythmic stimulation (after stim). Bottom: I-V plot of the steady-state average (±SD) normalized currents. P = 0.300, n = 5. C, top and middle: IK (top) elicited by a test pulse from –40 to 0 mV (middle), before (control) and after 40 min of rhythmic stimulation in (and washout of) 200 µM Cd2+ (Stim in Cd2+). Bottom: I-V plot of the steady-state average (±SD) and normalized currents. P = 0.618, n = 5.