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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
Neurotoxicology. Author manuscript; available in PMC 2010 September 1.
Published in final edited form as:
PMCID: PMC2796506
NIHMSID: NIHMS154995

Chronic lead exposure alters presynaptic calcium regulation and synaptic facilitation in Drosophila larvae

Abstract

Prolonged exposure to inorganic lead (Pb2+) during development has been shown to influence activity-dependent synaptic plasticity in the mammalian brain, possibly by altering the regulation of intracellular Ca2+ concentration ([Ca2+]i). To explore this possibility, we studied the effect of Pb2+ exposure on [Ca2+]i regulation and synaptic facilitation at the neuromuscular junction of larval Drosophila. Wild-type Drosophila (CS) were raised from egg stages through the third larval instar in media containing either 0, 100 μM or 250 μM Pb2+ and identified motor terminals were examined in late third-instar larvae. To compare resting [Ca2+]i and the changes in [Ca2+]i produced by impulse activity, the motor terminals were loaded with a Ca2+ indicator, either Oregon Green 488 BAPTA-1 (OGB-1) or fura-2 conjugated to a dextran. We found that rearing in Pb2+ did not significantly change the resting [Ca2+]i nor the Ca2+ transient produced in synaptic boutons by single action potentials (APs); however, the Ca2+ transients produced by 10 and 20 Hz AP trains were larger in Pb2+-exposed boutons and decayed more slowly. For larvae raised in 250 μM Pb2+, the increase in [Ca2+]i during an AP train (20 Hz) was 29% greater than in control larvae and the [Ca2+]i decay τ was 69% greater. These differences appear to result from reduced activity of the plasma membrane Ca2+ ATPase (PMCA), which extrudes Ca2+ from these synaptic terminals. These findings are consistent with studies in mammals showing a Pb2+-dependent reduction in PMCA activity. We also observed a Pb2+-dependent enhancement of synaptic facilitation at these larval neuromuscular synapses. Facilitation of EPSP amplitude during AP trains (20 Hz) was 55% greater in Pb2+-reared larvae than in controls. These results showed that Pb2+ exposure produced changes in the regulation of [Ca2+]i during impulse activity, which could affect various aspects of nervous system development. At the mature synapse, this altered [Ca2+]i regulation produced changes in synaptic facilitation that are likely to influence the function of neural networks.

Keywords: Pb2+, synapse, Drosophila, larvae, calcium, facilitation

Introduction

Chronic Pb2+ exposure can produce changes in the structure and function of the mammalian brain (Costa et al., 2004; Toscano and Guilarte, 2005). This is particularly true for synapses where Pb2+ exposure during prenatal and/or postnatal development altered paired-pulse facilitation and long-term potentiation in the rat hippocampus (Altmann et al., 1993; Gilbert et al., 1996; Gilbert and Mack, 1998; Lasley et al., 1993; Ruan et al., 1998). In addition, Pb2+ exposure during postnatal development (10 mg/ml Pb acetate in fluids or 4% Pb carbonate in food) has been shown to alter the density of dendritic spines and presumably synapses in the brains of rats and cats (Kiraly and Jones, 1982; Petit and LeBoutillier, 1979). The effects of Pb2+ exposure on synaptic development and plasticity could result from alterations in [Ca2+]i regulation. Intracellular Ca2+ can influence multiple steps in synaptic development; e.g., growth cone guidance (Jin et al., 2005), synapse formation (Xu et al., 2009) and synapse elimination and stabilization (Lohmann and Bonhoeffer, 2008; Pratt et al., 2003). At the mature synapse, altered [Ca2+]i regulation can influence long and short-term forms of synaptic plasticity (MacDonald et al., 2006; Zucker and Regehr, 2002).

The effects of Pb2+ on Ca2+ influx and efflux are well documented. Acute exposure to Pb2+ (nanomolar to micromolar) blocks Ca2+ influx through both invertebrate (Audesirk and Audesirk, 1989; Busselberg et al., 1990) and mammalian voltage-dependent Ca2+ channels (Audesirk and Audesirk, 1991; Audesirk and Audesirk, 1993; Busselberg et al., 1993; Evans et al., 1991; Peng et al., 2002). Also, micromolar concentrations of Pb2+ can inhibit the extrusion of Ca2+ by the plasma membrane Ca2+ ATPase (PMCA) in humans and rats (Bettaiya et al., 1996; Mas-Oliva, 1989; Sandhir and Gill, 1994) although the PMCA is stimulated by lower (nanomolar) Pb2+ concentrations (Ferguson et al., 2000; Mas-Oliva, 1989). However, it is not known whether the effects of Pb2+ exposure on synaptic development and plasticity are due to altered regulation of [Ca2+]i. In fact, the effect of Pb2+ exposure on the regulation of [Ca2+]i has not been well characterized. For example, there are no studies examining the effect of Pb2+ exposure on the changes in [Ca2+]i produced by impulse activity. It is difficult to examine the acute effects of Pb2+ on [Ca2+]i regulation since Pb2+ can enter through voltage-dependent Ca2+ channels (Simons and Pocock, 1987; Tomsig and Suszkiw, 1991) and Ca2+ indicators can respond to both Ca2+ and Pb2+ (Kerper and Hinkle, 1997; Tomsig and Suszkiw, 1990). However, the long-term effects of chronic Pb2+ exposure on [Ca2+]i can be measured in Pb2+-free saline. In fact, fura-2 has been used to measure resting [Ca2+]i in synaptosomes isolated from rats after postnatal in vivo exposure to Pb2+ (Sandhir and Gill, 1994).

In this study, we addressed the following unanswered questions. First, what is the effect of Pb2+ exposure during development on the presynaptic Ca2+ signals produced by impulse activity? Second, if there are changes in presynaptic [Ca2+]i regulation, are they correlated with changes in synaptic plasticity? To answer these questions we used the Drosophila larval neuromuscular junction (NMJ) one of the preferred systems for studies of the molecular basis for synaptic function, development and plasticity (Keshishian et al., 1996). In addition to its advantages for genetic analyses, the larval motor terminals are identifiable and accessible for electrophysiological and optical studies. Previously, Pb2+ exposure was shown to affect the development of behavior in Drosophila (Hirsch et al., 2003; Hirsch et al., 2009) and synaptic development at the larval NMJ (Morley et al., 2003).

Materials and Methods

To produce larvae, we added 20 female and male adult flies (wild type, Canton S) to 25 mm vials containing Instant Drosophila Medium Formula 4–24 (Carolina Biological Supply Company, Burlington, NC) mixed with either dH2O (nominal 0 Pb2+), 100 μM Pb acetate in dH2O or 250 μM Pb acetate in dH2O. Females thus laid eggs in control or leaded medium; once eggs hatched the larvae were raised in the same medium until the end of the third larval instar. At this developmental stage larvae leave the medium. We collected only female wandering 3rd instar larvae, identified by their gonads, for Ca2+ measurements and synaptic physiology. The doses used were based on previous work showing that 100 μM Pb acetate in dH2O affected development of the larval motor terminals (Morley et al., 2003), and 250 μM Pb acetate in dH2O influenced adult fly locomotion (Hirsch et al., 2009).

Loading the Ca2+ indicator

To measure [Ca2+]i, we used the dual-wavelength Ca2+ indicator fura-2 (Kd 594 nM) or the single-wavelength indicator, Oregon Green 488 BAPTA-1 (OGB-1; Kd 1180 nM) In both cases, they were coupled to 10,000 MW dextrans (Invitrogen, Carlsbad, CA). As in our previous studies (He et al., 2009; Lnenicka et al., 2006a), we loaded the indicator into the motor terminals using the technique developed by Macleod et al., 2002. Briefly, larvae were dissected in Schneider’s media (Sigma, St. Louis, MO) and the nerve supplying the third segment was cut. The cut end of the nerve was immediately sucked up into a snug-fitting pipette and then a small volume of indicator (5 mM) was introduced into the end of the pipette tip using very thin tubing. After 40 min, the nerve was removed from the pipette and the preparation was left at room temperature for another hour. The Schneider’s solution was then replaced with HL3 saline (Stewart et al., 1994) containing 1 mM Ca2+ for physiological measurements. For some experiments, the membrane-permeant, heavy-metal chelator N,N,N′,N′-tetrakis-(2-pyridylmethyl)ethylenediamine (TPEN; Invitrogen) was dissolved in ethanol (200 mM) and added to the saline for a final concentration of 100 μM.

Measurement of [Ca2+]i changes

The motor terminals were imaged using an upright, fixed-stage BH2 microscope (Olympus) equipped with epifluorescence, differential-interference-contrast optics, a water-immersion 40x Zeiss lens (NA 0.75) and a digital cooled-CCD camera (CoolSNAP HQ, Photometrics, Tucson, AZ). Excitation illumination from a 75W xenon arc lamp was passed through a Lambda-10 Optical Filter Changer (Sutter Instrument Co., Novato, CA). Fura-2 was excited using 360 ±5 and 380 ±5 nm bandpass filters (Chroma Technology Corp., Brattleboro, VT); excitation and emission wavelengths were separated with a 410 nm dichroic mirror and the emitted light was passed through a barrier filter of 510 ±10 nm. For OGB-1, we used a 480±15nm excitation filter, a 500 nm dichroic mirror and a high-pass 515 nm barrier filter. Images were captured using 100 msec exposures for fura-2. For OGB-1, images were streamed using 20 msec exposures (image size- 50 × 350 pixels) for single APs and 50 msec (image size- 100 × 350 pixels) exposures for AP trains. Metafluor 6.1 software (Molecular Devices, Downingtown, PA) was used for image acquisition and to measure fluorescent intensity at synaptic boutons and background fluorescence from the muscle. Metamorph 6.1 software (Molecular Devices) was used to measure bouton widths.

As in a previous study, we estimated the [OGB-1] in the synaptic terminals by measuring the fluorescence intensity for a range of OGB-1 concentrations in capillary tubes with an inside diameter of approximately 20 μm (He et al., 2009). The fluorescent values were normalized to a 20 μm diameter giving a linear plot of [OGB-1] versus fluorescent intensity. For each terminal, the average resting fluorescence was measured at a region of uniform terminal width (apparent diameter) and the fluorescence intensity was normalized to a 20 μm diameter. These capillary tubes contained 40 nM Ca2+; we assumed a resting [Ca2+]i of 40 nM in the motor terminals (Klose et al., 2008). This fluorescent value was then converted to [OGB-1] using the above plot. Capillary tubes with a 20 μm inside diameter were produced from 50 μm capillary tubes (VitroCom, Mountain Lakes, NJ) using a Narishige PN-30 electrode puller (Narishige International USA Inc., East Meadow, NY).

Synaptic physiology

The nerve was stimulated with a suction electrode connected to a S11 stimulator (Grass-Telefactor, West Warwick, RI) and excitatory postsynaptic potentials (EPSPs) were recorded from muscle fibers with sharp electrodes using an Axoclamp 2A amplifier (Axon Instruments Inc., Foster City, CA). pCLAMP 9.2 software and a Digidata 1200A digitizer (Axon Instruments Inc., Foster City, CA) were used to acquire and measure EPSPs.

Data analysis

Sigmaplot 10.0 and SPSS 16.0 (SPSS Inc. Plover, WI) were used for data transformation and statistical analysis. The background fluorescence was subtracted from the bouton fluorescence for each image. For fura-2, we calculated the ratio of fluorescence produced by 360 and 380 nm excitation (360/380). For OGB-1, the percentage change in fluorescence (ΔF/F) was calculated by 100 × (fluorescence − initial fluorescence)/initial fluorescence. Single exponentials were fit to the ΔF/F decay to determine the [Ca2+]i decay time constant (τ). We only report [Ca2+]i decay τ where the fit to a single exponential gave r2 > 0.9 in order to eliminate noisy measurements. The increase in [Ca2+]i during trains of impulses was measured after the [Ca2+]i increase reached a plateau by averaging the last 20 measurements of the train. Statistical analyses were performed using t-tests with a Welch correction for unequal variances or an ANOVA followed by a post-hoc analysis (Winer B.J., 1962); p values were adjusted using either a Bonferroni correction when variances were not significantly different or a Games-Howell correction if variances were significantly different (Kirk R.E., 1995). All n values were reported as (# boutons, # animals) unless otherwise noted.

Results

To examine the developmental effect of prolonged Pb2+ exposure on intracellular [Ca2+]i regulation at synaptic terminals, Drosophila larvae were raised from egg stages to the end of the third-instar larval stage in either control media (nominal 0 Pb2+ concentration) or in leaded media containing 100 μM or 250 μM Pb2+. In a previous study, the Pb2+ burdens were measured in larvae raised in 100 μM Pb2+ using inductively-coupled mass spectrometry; this gave values of 12.37 ng Pb2+ per larva, which had an average wet weight of 1.6 mg (Morley et al., 2003). The larval mid and hindgut contain about 73% of the Pb2+ (Wilson, 2004) and the hemolymph is approximately 40% of wet body weight for insect larvae (Nation, 2001). Thus, the Pb2+ concentration in the hemolymph would be 25 μM if all of the Pb2+ were in the hemolymph. However, the Pb2+ concentration is presumably much less than this since Pb2+ would also be distributed in the intracellular fluids and bound to cellular components. If fact, the hemolymph makes up only half of the total larval water content since about 80% of wet larval weight is due to water (Folk 2001). In conclusion, it appears likely that the levels of total Pb2+ in the larval hemolymph are micromolar, which falls within the range of Pb2+ blood levels observed to influence nervous system development and synaptic plasticity in other organisms (Cline et al., 1996; Gilbert et al., 1996; Lorton and Anderson, 1986; Petit and LeBoutillier, 1979).

In wandering third-instar larvae, we compared [Ca2+]i in the Ib motor terminals innervating muscle fibers 6 and 7 (MF 6/7). MF 6/7 are innervated by Is and Ib motor terminals supplied by two separate motor neurons (Hoang and Chiba, 2001; Lnenicka and Keshishian, 2000). The terminals can be distinguished by their different morphology: Ib terminals have large synaptic boutons and Is terminals have small boutons (Kurdyak et al., 1994).

Increase in [Ca2+]i produced by impulse activity

The increase in [Ca2+]i produced by single APs and trains of APs was examined in control and Pb2+-reared larvae. We used the single-wavelength Ca2+ indicator OGB-1, which allowed us to measure the rapid, “volume-averaged” changes in [Ca2+]i produced by single APs (Lnenicka et al., 2006a). The change in OGB-1 fluorescence (ΔF/F) produced by single APs (ΔF/FAP) and AP trains (ΔF/Ftrain) was determined for individual synaptic boutons (Fig 1). In general, the Pb2+-exposed synaptic boutons showed similar ΔF/FAP amplitudes but larger ΔF/Ftrain amplitudes than boutons from larvae grown in 0 Pb2+.

Figure 1
Ca2+ transients recorded from synaptic boutons in control and Pb2+-exposed animals. Left: Ib terminals on MF 6/7 filled with OGB-1 from control larvae and those exposed to Pb2+. The arrows point to two typical synaptic boutons where Ca2+ transients were ...

The differences in Ca2+ transients for control and Pb2+-exposed larvae were quantified by comparing a number of animals. In these experiments, it was necessary to use a similar [OGB-1] in the terminals of control and experimental larvae; the Ca2+ indicator acts as a buffer and high concentrations will decrease the amplitude of the ΔF/FAP and reduce the rate of [Ca2+]i decay at the end of the Ca2+ transient (Neher, 1995). In two experiments, the [OGB-1] was very high (> 100 μM) and these experiments were not included. For the remaining experiments, the [OGB-1] was similar for terminals from control (43.3 ±7.3 μM, 11 terminals), 100 μM Pb2+-exposed (43.3 ±8.0 μM, 7 terminals) and 250 μM Pb2+-exposed larvae (49.2 ±4.1 μM, 7 terminals). In addition, the Ca2+ transients can be influenced by bouton size so it was necessary to compare bouton populations that were of similar size (Lnenicka et al., 2006a); bouton width was similar for control (2.5 ±0.1 μm; 94, 11), 100 μM Pb2+ (2.6 ±0.1 μm; 59, 7) and 250 μM Pb2+ terminals (2.7 ±0.1 μm; 121, 7).

The ΔF/FAP amplitude was not significantly different for Pb2+-reared larvae compared to controls; however, Pb2+-reared larvae showed a significantly larger ΔF/Ftrain amplitude than controls (Fig. 2). The ΔF/Ftrain amplitude increased with increasing concentrations of Pb2+. For 20 Hz stimulation, the ΔF/Ftrain amplitude was 16% and 29% larger than controls for 100 μM and 250 μM Pb2+, respectively. An increase in the [Ca2+]i plateau during an AP train can be produced by greater Ca2+ influx or a reduction in the rate of Ca2+ extrusion (Tank et al., 1995). Since the ΔF/FAP amplitudes were similar for control and Pb2+-exposed terminals, it is unlikely that the greater ΔF/Ftrain amplitude resulted from differences in Ca2+ influx. Rather, this difference in amplitude likely reflected differences in Ca2+ extrusion.

Figure 2
The Ca2+ transient amplitude in synaptic boutons from control larvae and those exposed to 100 or 250 μM Pb2+. The OGB-1 ΔF/F was measured at the peak of the Ca2+ transient for single APs (ΔF/FAP peak) and at the plateau for AP ...

The decay of [Ca2+]i after an AP train reflects the rate of Ca2+ extrusion: a higher extrusion rate will result in a smaller [Ca2+]i decay time constant (τdecay). We compared the [Ca2+]i τdecay for control and Pb2+ exposed terminals after single APs and AP trains. There was no clear effect of Pb2+ exposure on the ΔF/FAP τdecay since the τdecay was significantly greater for larvae exposed to 100 μM Pb2+, but not for those exposed to 250 μM Pb2+ (Fig. 3). For AP trains, Pb2+ exposure produced an increase in the τdecay in a dose-dependent manner (Fig. 3, ,4).4). For 20 Hz stimulation, the ΔF/Ftrain τdecay was 27% and 69% greater than controls for animals exposed to 100 μM and 250 μM Pb2+, respectively.

Figure 3
The Ca2+ transient decay time constant in synaptic boutons from control and Pb2+-exposed animals. The τdecay of the ΔF/F was measured after a single AP (ΔF/FAP τdecay) or at the end of an AP train (ΔF/Ftrain τ ...
Figure 4
Ca2+ transient decay after 20 Hz AP train normalized to the final response at the end of the AP train. The mean normalized ΔF/Ftrain decay was compared for animals exposed to 0 μM Pb2+ (68, 10), 100 μM Pb2+ (53,7) and 250 μM ...

It was possible that the Pb2+-exposed terminals still contained free Pb2+ that bound to OGB-1 and compromised our [Ca2+]i measurements. To test for this, we added 100 μM TPEN, a membrane-permeant, heavy-metal chelator; TPEN has a much higher affinity for Pb2+ than Ca2+ and reduces intracellular [Pb2+]i, but not resting [Ca2+]i (Arslan et al., 1985). In 8 larvae raised in 250 μM Pb2+, we measured resting OGB-1 fluorescence and ΔF/F in the same boutons before and after applying TPEN. We found that resting fluorescence (25.1 ±1.4AU) and ΔF/FAP (8.2 ±0.5 %) did not significantly change after TPEN addition (26.4 ±1.5 AU and 8.2 ±0.6 %, respectively; p> 0.10). Note that the ΔF/FAP amplitude was small because we included terminals containing high concentrations of OGB-1. This confirms that OGB-1 can be used to accurately measure [Ca2+]i changes in Pb2+-exposed terminals.

Comparison of resting [Ca2+]i

Pb2+ exposure has been shown to produce both an increase and decrease in resting [Ca2+]i in neurons (Ferguson et al., 2000; Sandhir and Gill, 1994). We performed ratiometric Ca2+ measurements using fura-2 to compare resting [Ca2+]i in synaptic terminals from control and Pb2+-exposed larvae. The fura-2 ratio for experimental and control terminals was also compared in the presence of TPEN in case there was any free Pb2+ remaining in the Pb2+-exposed terminals; fura-2 binds Pb2+ with a high affinity and this can produce an increase in the fura-2 ratio (Tomsig and Suszkiw, 1990). The change in indo-1 or fura-2 fluorescence produced by an increase in [Pb2+]i was shown to be reversed by the addition of TPEN (Kerper and Hinkle, 1997; Mazzolini et al., 2001).

We found no significant difference in the resting fura-2 ratios for 0 Pb2+ (0.35 ±0.01) and 250 μM Pb2+ terminals (0.33 ±0.02) before adding TPEN (p > 0.10). After adding TPEN, there was also no significant difference in the fura-2 ratios for 0 Pb2+ (0.36 ±0.01) and 250 μM Pb2+ terminals (0.32 ±0.02; p > 0.10). Also, the fura-2 ratio in the Pb2+-exposed terminals was not significantly reduced after adding TPEN (p> 0.10). The data were from 4 control and 4 Pb2+-exposed animals and an ANOVA was used for statistical comparisons.

Synaptic facilitation

Synaptic facilitation is dependent on the “residual” Ca2+ that equilibrates in the terminal after entering through voltage-dependent Ca2+ channels (Zucker and Regehr 2002). Since we have found that an AP train results in a greater buildup of residual Ca2+ in Pb2+-exposed terminals, these terminals might show greater synaptic facilitation compared to control terminals. To test this, we measured EPSP amplitude during low-frequency stimulation and during trains of APs delivered at higher frequencies. By performing these experiments on muscle fiber 5, which is innervated by a single axon, we could measure facilitation at a single synapse (Lnenicka et al., 2006b).

We compared synaptic physiology in control larvae and those raised in 250 μM Pb2+ since this concentration produced a greater effect on [Ca2+]i regulation than 100 μM Pb2+. To determine the EPSP amplitude for single APs, the axon was stimulated at 0.5 Hz, a stimulation frequency that did not produce facilitation. To compare synaptic facilitation, we stimulated the axon at 20 Hz; we previously showed that Pb2+-exposed and control terminals showed clear differences in the buildup of residual Ca2+ at this frequency. We found that there was greater synaptic facilitation in Pb2+-exposed animals during 20 Hz stimulation compared to control animals (Fig. 5). We found that the amplitude of the EPSP produced by single APs was not significantly different in 0 Pb2+ and 250 μM Pb2+ larvae (Fig. 6). However, the EPSP amplitude during 20 Hz stimulation was significantly greater for Pb2+-exposed larvae compared to controls.

Figure 5
EPSPs recorded from muscle fibers in control and Pb2+-exposed larvae during 20 Hz stimulation. Top: Typical EPSPs recorded from muscle fiber 5 during 20 Hz stimulation. The EPSPs were larger in larvae raised in media containing 250 μM Pb2+ compared ...
Figure 6
Mean EPSP amplitudes and synaptic facilitation in control larvae and those exposed to 250 μM Pb2+. Left: The EPSP amplitude produced by single APs (0.5 Hz) was not significantly different in 0 Pb2+ and 250 μM Pb2+ larvae. During 20 Hz ...

The differences in the EPSP amplitude seen during 20 Hz stimulation appear to be mainly due to differences in the buildup of synaptic facilitation. During a train of APs, the enhancement of EPSP amplitude can result from the buildup of facilitation plus the onset of the longer-lasting augmentation and post-tetanic potentiation (PTP). Synaptic facilitation has a decay time constant of 10s to 100s of milliseconds, augmentation decays with a time constant of 5–10 sec. and PTP has a decay time constant of 30 sec to several minutes (Magleby and Zengel, 1976; Magleby and Zengel, 1975; Mallart and Martin, 1967). The differences in EPSP amplitude for Pb2+-exposed and control larvae were nearly gone 2 seconds after the end of the 20 Hz train (Fig. 5). Thus, the differences between Pb2+-exposed and control terminals are only seen during synaptic facilitation and not during the augmentation and PTP phases. Overall, the synaptic facilitation during the 20 Hz train was significantly greater in Pb2+-exposed larvae than in control larvae (Fig. 6).

Discussion

Pb2+ exposure influences the [Ca2+]i increase produced by APs

We found that the Ca2+ transient produced by single APs was not influenced by chronic Pb2+ exposure; however, Pb2+ exposure resulted in a larger Ca2+ transient during AP trains. The ΔF/FAP amplitude was similar in control synaptic boutons and those exposed to 100 or 250 μM Pb2+. Since the amplitude of the single-AP Ca2+ transient is determined by the amount of Ca2+ influx and the concentration of fast Ca2+ buffers, it appears that Ca2+ influx was similar in control and Pb2+-exposed terminals. Alternatively, there could have been Pb2+-related changes in Ca2+ influx that were counterbalanced by changes in fast Ca2+ buffering. This seems unlikely since changes in Ca2+ buffering would result in a change in the ΔF/FAP τdecay (Neher, 1995) and we did not see a consistent effect of Pb2+ exposure on the ΔF/FAP τdecay. The inhibition of Ca2+ influx by acute Pb2+ exposure (nanomolar to micromolar) has been demonstrated for a variety of voltage-dependent Ca2+ channels in both invertebrate (Audesirk and Audesirk, 1989; Busselberg et al., 1990) and mammalian voltage-dependent Ca2+ channels (Audesirk and Audesirk, 1991; Audesirk and Audesirk, 1993; Busselberg et al., 1993; Evans et al., 1991; Peng et al., 2002). In most cases, the inhibition of Ca2+ influx by Pb2+ was reversible (Audesirk, 1993) and we assume that any inhibitory effects of Pb2+ were washed out before we performed our measurements. Thus, our measurements do not rule out the possibility that Pb2+ in the hemolymph inhibited Ca2+ influx in vivo.

For AP trains, chronic Pb2+ exposure resulted in a linear dose-dependent increase in the ΔF/Ftrain amplitude and τdecay. We assume that both of these effects resulted from a decrease in the rate of Ca2+ extrusion by the PMCA. During an AP train, the buildup of [Ca2+]i results in greater Ca2+ extrusion until Ca2+ influx and efflux per unit time are equal and the plateau is reached (Tank et al., 1995); a reduction in the Ca2+ extrusion rate constant will result in a greater ΔF/Ftrain amplitude. At the end of the train, the [Ca2+]i decay reflects the rate of Ca2+ extrusion and a lower extrusion rate will result in a greater ΔF/Ftrain τdecay. The PMCA is responsible for Ca2+ extrusion from these larval synaptic terminals; inhibition of the PMCA resulted in an increase in ΔF/Ftrain amplitude and the ΔF/Ftrain τdecay and immunostaining showed localization of the PMCA at the NMJ (Lnenicka et al., 2006a). It appears unlikely that Pb2+ also acts on other Ca2+-clearance mechanisms to produce the increase in ΔF/Ftrain amplitude since blocking Ca2+ extrusion by the Na/Ca exchanger or Ca2+ uptake by mitochondria or ER did not result in an increase in ΔF/Ftrain amplitude in a previous study (Lnenicka et al., 2006a).

There was no clear and consistent effect of Pb2+ exposure on the ΔF/FAP τdecay (the ΔF/FAP τdecay increased significantly for the 100 μM Pb2+ group and then dropped for the μM Pb2+ group). This may be because the PMCA plays a greater role in determining the ΔF/Ftrain τdecay than the ΔF/FAP τdecay. This is supported by the finding that blocking the PMCA had a greater effect on ΔF/Ftrain τdecay than on ΔF/FAP τdecay (Lnenicka et al., 2006a). Factors other than the PMCA may play a role in the ΔF/FAP decay; e.g., studies of crayfish motor terminals indicate that slow Ca2+ buffers play a role in the ΔF/FAP decay (Lin et al., 2005).

Pb2+ exposure has been shown to influence PMCA activity in other systems. Acute exposure to micromolar concentrations of Pb2+ reduced the activity of the PMCA in humans and rats (Bettaiya et al., 1996; Mas-Oliva, 1989; Sandhir and Gill, 1994). The in vitro studies indicate that Pb2+ inhibits PMCA activity by binding directly to the PMCA (Mas-Oliva, 1989; Sandhir and Gill, 1994). Studies examining the effect of in vivo chronic Pb2+ application have shown a persistent inhibition of PMCA activity; i.e., PMCA activity was inhibited when measured in nominal Pb2+-free solutions. For example, the PMCA activity in erythrocytes from mothers and newborns was negatively correlated with hair Pb2+ levels (Campagna et al., 2000). In addition, rats exposed to Pb2+ intragastrically (50 mg/kg body weight) for 8 weeks showed a 31% decrease in PMCA activity per total protein for synaptic plasma membranes isolated from the brain (Sandhir and Gill, 1994). Our results also suggest a persistent effect of Pb2+ on PMCA activity and our experiments using TPEN showed no evidence for free Pb2+ remaining in the synaptic terminals. The persistent effects of chronic Pb2+ exposure could be due to irreversible binding of Pb2+ to the PMCA. Alternatively, Pb2+ exposure could have reduced the expression of PMCA. The level of [Ca2+]i is one factor regulating PMCA expression (Guerini et al., 1999) and it may be that Pb2+ exposure decreased PMCA expression by reducing Ca2+ influx through Ca2+ channels.

In mammals, there are 4 PMCA genes and over 20 possible splice variants (Strehler and Treiman, 2004; Strehler and Zacharias, 2001) expressed in a tissue- and development-specific manner; all cell types express at least one of the isoforms (Strehler and Treiman, 2004). Nervous tissue appears to show the greatest abundance and variety of PMCA expression: many of the isoforms have been demonstrated in the brain (Strehler and Treiman, 2004). Drosophila has fewer PMCA genes and isoforms than found in mammals. Based upon the annotation of the genome, and BLASTp analysis of a mouse PMCA (PMCA-4) against the Drosophila annotated protein sequences, Drosophila has only a single highly homologous PMCA gene (CG42314) and 6 potential isoforms (E value = 0).

We found no effect of chronic Pb2+ exposure on resting [Ca2+]i and this appears unexpected since the PMCA plays an important role in maintaining resting [Ca2+]i. However, previous in vitro studies have reported an increase in resting [Ca2+]i during exposure to 5 μM Pb2+ (Schanne et al., 1989) and a decrease in resting [Ca2+]i (Ferguson et al., 2000) during exposure to 0.1 μM Pb2+. This apparently occurs because Pb2+ can stimulate PMCA activity as well as inhibit it. Normally, an increase in [Ca2+]i activates the PMCA through the formation of Ca-calmodulin, which binds to the C-terminal tail of the PMCA releasing it from autoinhibition (Strehler and Treiman 2004). Pb2+ can substitute for Ca2+ in calmodulin activation (Goldstein and Ar, 1983; Habermann et al., 1983; Kern et al., 2000) and this can result in an increase in PMCA activity (Ferguson et al., 2000). Synaptosomes isolated from rats exposed to Pb2+ in vivo (50 mg/kg body weight) showed an increase in resting [Ca2+]i due to reduced PMCA activity even though there was also an increase calmodulin activity (Sandir and Gill, 1994). The balance of PMCA inhibition and activation likely determines the effect of Pb2+ on resting [Ca2+]i; this likely depends on the Pb2+ concentration since activation of calmodulin occurs at nanomolar concentrations and PMCA inhibition at micromolar concentrations (Ferguson et al., 2000). In our studies, there may have been an equal balance of PMCA inhibition and excitation during resting [Ca2+]i; however, the effects of PMCA inhibition may have predominated at higher [Ca2+]i. For example assuming Michaelis-Menton kinetics and fewer functional PMCAs (lower Vmax) in Pb2+-exposed larvae, the effects of PMCA inhibition on the steady-state [Ca2+]i would be accentuated at a higher [Ca2+]i. As Ca2+ influx increased during AP trains, the steady-state [Ca2+]i concentration would increase more rapidly in Pb2+-exposed animals since the PMCA would approach saturation at a lower [Ca2+]i.

Synaptic facilitation

We tested whether the change in [Ca2+]i regulation resulting from Pb2+ exposure produced changes in synaptic plasticity. We found that during a train of impulses there was a greater increase in EPSP amplitude for Pb2+-exposed terminals than for control terminals. This activity-dependent synaptic enhancement is the most common form of synaptic plasticity and is found at most, if not all synapses (Zucker and Regehr, 2002). It results from an increase in transmitter release and can be divided into synaptic facilitation, augmentation and PTP based upon differences in time course. It appeared that differences in synaptic enhancement seen for control and Pb2+-exposed terminals were due to differences in the buildup of synaptic facilitation.

The greater synaptic facilitation seen at the Pb2+-exposed terminals is likely due to the greater increase in [Ca2+]i seen at these terminals. Although the mechanisms underlying synaptic enhancement are not understood, studies of synapses in vertebrates and invertebrates have shown that facilitation, augmentation and PTP appear to be dependent upon residual Ca2+ (Zucker and Regehr, 2002). For example, adding the Ca2+ chelator ethylene glycol tetraacetic acid (EGTA) alters the buildup and decay of Ca2+ and synaptic enhancement during a train at mammalian central synapses and the lobster NMJ (Ogawa et al., 2000; Regehr et al., 1994).

Previous studies examining the effects of in vivo Pb2+ exposure on synaptic plasticity have mainly focused on long-term potentiation (LTP). Postnatal Pb2+exposure increased the threshold for producing LTP, decreased its amplitude and reduced the duration of LTP in the rat hippocampus, possibly due to the effect of Pb2+ on the NMDA receptor (Gilbert et al., 1996; Gilbert and Mack, 1998; Lasley et al., 1993; Ruan et al., 1998). However, the effect of Pb2+ exposure on short-term plasticity may differ since exposure to levels of Pb2+ that impaired LTP resulted in an increase in the amplitude of short-term potentiation (STP) in the rat hippocampus (Grover and Frye, 1996). Since STP may involve presynaptic mechanisms (Lauri et al., 2007), the increase in STP could involve an increase in residual Ca2+ as seen at the larval NMJ.

The changes that we observe in [Ca2+]i regulation and synaptic facilitation at the neuromuscular synapses are likely to occur at synapses in the central nervous system since the PMCA plays an important role in Ca2+ extrusion from synaptic terminals in the central nervous (Juhaszova et al., 2000) (Morgans et al., 1998) and synaptic facilitation at central synapses is dependent on residual Ca2+ (Atluri and Regehr, 1996). In fact, reduced PMCA activity has been shown to enhance synaptic facilitation at central synapses; these studies involved blocking PMCA activity pharmacologically or knocking out expression of one of the PMCA isoforms (Empson et al., 2007; Jensen et al., 2007).

Acknowledgments

This work was supported by the Environmental Health Sciences Center in Molecular and Cellular Toxicology with Human Applications Grant P30 ES06639 at Wayne State University, NIH R01 grant ES012933 (D.M.R.) and NSF grant IOB 0543835 (G.A.L.).

Footnotes

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