INTRODUCTION

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

In the mammalian olfactory system, odors are encoded by the differential activation of a large multigene family of olfactory receptors expressed in different subsets of olfactory receptor cells (Buck and Axel 1991; Malnic et al. 1999). Cells expressing the same receptor project to a small subset of glomeruli within a large glomerular array on the surface of the olfactory bulb, creating a stimulus-specific two-dimensional spatial representation of olfactory receptor activation (Mombaerts et al. 1996; Wang et al. 1998). This pattern of glomerular activity is relayed to layers of projection neurons, the mitral and tufted cells (Price and Powell 1970a). A mitral cell receives glomerular synaptic input via the distal tuft of a primary (=apical) dendrite extending vertically from its soma. The mitral soma also radiates secondary (=basal or lateral) dendrites which extend horizontally ~1,000 µm across the external plexiform layer of the bulb (Mori et al. 1983; Orona et al. 1984). These dendrites are linked laterally by an extensive network of reciprocal dendrodendritic synaptic connections with granule cells (Jackowski et al. 1978; Rall et al. 1966). The lateral connections mediate excitatory-inhibitory interactions, and the current view is that they can shape both the spatial and temporal patterns of mitral cell activity that are thought to encode the intensity and quality of odors (Laurent 1999).

Mitral cell activity is controlled by a complex interplay between intrinsic conductances and synaptic inputs. Excitatory postsynaptic potentials (EPSPs) originating in the distal tuft initiate action potentials either in the soma or in the primary dendrite, depending on the level of somatic inhibition (Chen et al. 1997). The mitral cell membrane exhibits subthreshold bistability with a depolarized plateau potential (Heyward et al. 2001), and action-potential timing can lock to subthreshold membrane potential oscillations, which can be reset by inhibitory postsynaptic potentials (IPSPs) (Chen and Shepherd 1997; Desmaisons et al. 1999). Action potentials activate voltage-sensitive Ca²⁺ channels (Cinelli and Salzberg 1990, 1992; Mori et al. 1981; Wang et al. 1996), triggering glutamate release from the mitral cell at reciprocal synapses (Isaacson and Strowbridge 1998). The glutamate activates granule cell spines, which release GABA to inhibit the mitral cell (Isaacson and Strowbridge 1998; Jahr and Nicoll 1980; Nowycky et al. 1981; Rall et al. 1966). In addition to this negative feedback inhibition, mitral cells receive lateral inhibition from granule cells activated independently by other mitral cells (Isaacson and Strowbridge 1998; Margrie et al. 2001; Rall et al. 1966). This inhibitory circuitry is augmented by other types of GABAergic interneurons distinguished by parvalbumin immunoreactivity, which make dendrodendritic synapses with the mitral cell soma and primary dendritic shaft (Crespo et al. 2001; Toida et al. 1994, 1996). Glutamate released from mitral cells also activates glutamate autoreceptors on mitral cells, which can modulate burst firing (Friedman and Strowbridge 2000; Salin et al. 2001), and there is evidence for a positive feedback excitatory pathway between mitral cells and interneurons (Didier et al. 2001).

The effect of lateral inhibition on a mitral cell depends on the strength and location of the inhibitory input, and its impact on local signaling processes. Anatomical studies have demonstrated symmetric, presumably inhibitory synapses on the mitral cell membrane (Crespo et al. 2001; Price and Powell 1970a,b; Rall et al. 1966; Sassoe-Pognetto and Ottersen 2000; Toida et al. 1994, 1996). On the secondary dendrite, the ability of such synapses to block firing depends on their distance from the soma, the range of current shunting along the dendrite, and the local density of functional postsynaptic receptors. The range of lateral inhibition is of special interest because of its presumed role in shaping spatial activity patterns. The extensiveness of the secondary dendrites suggests that more distal dendritic elements may be electrotonically decoupled from the soma. Distal GABAergic inhibition could then regulate dendritic electrical signaling locally, independent of the soma. Mitral cell dendrites are presynaptic structures, and presynaptic GABA receptors are well known to modulate neurotransmitter release (Dudel and Kuffler 1961; Eccles et al. 1963; MacDermott et al. 1999; Nicoll and Alger 1979). Inhibition can alter the amplitudes and waveforms of action potentials invading presynaptic terminals (Baxter and Bittner 1991; Segev 1990; Zhang and Jackson 1995), which can strongly impact calcium influx and transmitter release (Sabatini and Regehr 1997). Invasion of mitral cell dendrites by backpropagating action potentials provides the depolarization required to initiate calcium influx for triggering dendritic glutamate release (Bischofberger and Jonas 1997; Isaacson and Strowbridge 1998; Margrie et al. 2001). Focal modulation of action potentials and calcium influx in the secondary dendrites by lateral inhibition has the potential to modify spatial patterns of dendrodendritic transmission across the bulb.

This paper describes the effect of localized inhibition on the initiation and backpropagation of action potentials in the secondary dendrites of rat mitral cells. Using laser flash photolysis of caged GABA, inhibition was applied locally to the soma and dendrites of mitral cells in olfactory bulb slices during somatic and dendritic whole cell recording. The advantage of this approach is that inhibition can be applied reproducibly with high spatial and temporal resolution (Katz and Dalva 1994; Wang and Augustine 1995). First, the conductance activated by uncaging GABA on the soma and dendrites of mitral cells was characterized. The magnitude of photoinhibition was then adjusted to approximately match physiological levels of inhibition inferred from inhibitory postsynaptic currents (IPSCs) from presynaptic spines of interneurons. This laid the groundwork for applying the method to probe the range of lateral inhibition and study the effect of inhibition on backpropagating action potentials.

	METHODS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Slice preparation

Horizontal olfactory bulb slices (350-µm thick) were prepared from 21- to 28-day-old male CD rats (Charles River). Animals were killed by overdose of halothane anesthesia (saturated vapor), and the olfactory bulbs were removed immediately into ice-cold sucrose artificial cerebrospinal fluid (ACSF) containing (in mM) 240 sucrose, 2.5 KCl, 10 Na-HEPES, 10 D-glucose, 1 CaCl₂, 4 MgCl₂, and 0.2 ascorbic acid, pH 7.2 with HCl, 317 mOsm, bubbled continuously with oxygen. Slices were cut in ice-cold sucrose ACSF with a vibrating razor blade (60 Hz) and allowed to recover for 1-3 h in an enclosed interface chamber containing high-Mg²⁺ ACSF (which was composed of, in mM, 124 NaCl, 2.5 KCl, 26 NaHCO₃, 1.25 NaH₂PO₄, 10 D-glucose, 1 CaCl₂, and 3 MgCl₂, 298 mOsm) bubbled continuously with 95% O₂-5% CO₂. A 20-gauge needle outlet allowed the gas to escape from the chamber under slight positive pressure. The recovery chamber was prewarmed to 30°C and left to cool slowly to room temperature (22°C). Slices were subsequently transferred to a second enclosed interface chamber containing standard ACSF (in mM): 124 NaCl, 2.5 KCl, 26 NaHCO₃, 1.25 NaH₂PO₄, 25 D-glucose, 2 CaCl₂, and 1.3 MgCl₂, 312 mOsm, bubbled with 95% O₂-5% CO₂, at 22°C. Slices remained in storage in the second chamber for 4 h before recording.

Electrophysiological recording

Slices were transferred to a small custom-designed Plexiglas chamber for submerged perfusion at 2 ml/min with standard ACSF at 25°C, bubbled with 95% O₂-5% CO₂. Temperature was regulated by a custom-built stage heater and an indium tin oxide heated coverslip (Cell MicroControls) forming the bottom of the recording well. Mitral cell somata and dendrites were visualized with a Nikon E600 FN upright microscope equipped with a Leica HCX APO L 63X/0.90 water-immersion objective, visible and infrared differential interference contrast (IR-DIC) optics, and an infrared video camera (C2400-79H, Hamamatsu Photonics K. K.). For whole cell recordings, voltage-clamp measurements were made with a CsCl pipette solution containing (in mM) 126.3 CsCl, 4.9 KCl, 25.2 K-HEPES, 0.2 K-EGTA, 1.9 Mg-ATP, 0.3 Na-GTP, 1 MgCl₂, 3.9 Na₂-phosphocreatine, and 6.3 biocytin, pH 7.2, 312 mOsm, E_Cl = 1.2 mV. Current-clamp measurements were made with a K-methylsulfate pipette solution containing (in mM): 123 K-CH₃SO₄, 4.7 KCl, 24.6 K-HEPES, 0.2 K-EGTA, 1.9 Mg-ATP, 0.3 Na-GTP, 0.9 MgCl₂, 3.8 Na₂-phosphocreatine, and 6.1 biocytin, pH 7.2, 312 mOsm, E_Cl = 59.8 mV. Pipette input resistance was 3-8 M for somatic recordings and 8-15 M for dendritic recordings. In some recordings, pharmacological agents were added to the bath: 1 µM TTX, 50 µM bicuculline methiodide (BMI), 60 µM 2-amino-5-phosphonopentanoic acid (AP-5), or 10 µM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX; all from Sigma RBI).

Whole cell voltage-clamp recordings were made with an EPC-8 patch-clamp amplifier (HEKA Electronics), and current-clamp recordings were made with two BVC-700 microelectrode amplifiers (Dagan) in bridge mode with electrode capacitance and series resistance compensation or an EPC-8 patch-clamp amplifier in fast current-clamp mode. Amplifiers were tested pairwise with dual recordings on the mitral cell soma to verify that there was no significant action potential distortion between the instruments. Spike waveforms recorded by two BVC-700 amplifiers were closely matched. With 100-pA current pulse injection, the ratio of spike amplitudes measured between the two was 0.996 ± 0.005 (n = 9 spikes) and the ratio of spike widths was 0.97 ± 0.01 (n = 9 spikes). Amplifiers were controlled through analog output boards driven by patch-clamp software written in LabVIEW (National Instruments). Data were acquired at 50-kHz, 16-bit resolution by two simultaneous-sampling dynamic signal analysis A/D boards (National Instruments), also controlled by software written in LabVIEW.

Laser flash photolysis

Once stable whole cell recordings were achieved, the perfusion system was switched to recycling mode (4.3 ml total volume of bubbled standard ACSF) to allow introduction of caged compounds and pharmacological agents. In this mode, a peristaltic pump removed ACSF from the chamber downstream of the slice and returned it upstream into an enclosed, elevated inlet channel with bubbling port; from there the solution drained by gravity back into the slice recording well. Caged neurotransmitter (O-CNB-caged GABA, at 270 µM or 400 µM; or -CNB-caged glutamate, at 320 µM; Molecular Probes) was added to the bath by injection into a loop manifold. Stock solutions of the caged compounds (6 or 9 mM) were made in standard ACSF at pH 4.00 to inhibit spontaneous hydrolysis, stored in the dark at 25°C, and thawed immediately before introduction into the recording chamber. Pharmacological agents were similarly injected. For focal flash photolysis of caged neurotransmitter, the beam from an Innova 90C argon ion laser (Coherent, Santa Clara, CA) was steered by mirrors into the fluorescence port of the Nikon E600. The output beam (351, 364 nm) was attenuated 40-fold by neutral density filters, reflected off a dichroic mirror, passed through the DIC prism, and focused by the Leica objective, whose apochromatic design allowed simultaneous IR imaging and UV photolysis in the same focal plane. The center of the beam in the focal plane was calibrated by imaging a fluorescent spot on a thin film of crystallized Lucifer yellow. The laser optics and microscope were moved over a fixed stage along x and y axes using an optical bench (TMC) driven by DC motors (Polytec PI), and the focal plane was positioned with a PIFOC piezoelectric translator (P-723, Polytec PI). Motion-control software was written in LabVIEW and integrated into the patch-clamp program. The uncaging beam was gated with an electronic shutter (Uniblitz, Vincent Associates), and the intensity of the output beam was set using the internal laser power meter. Measurements with a calibrated photodiode (United Detector Technology) verified a linear relationship between internal power meter reading and photocurrent. Approximate energy delivered to the cell per flash is quoted based on nominal attenuation factors of 0.65 for the objective and 0.90 for the DIC prism. In all experiments, the power of the unattenuated output beam was 50 mW. Flash duration for a given TTL pulse to the shutter driver was also calibrated using the photodiode (e.g., 1.24-ms flash for a 1-ms TTL pulse to the LS2 laser shutter), and timing delays for shutter opening were compensated in the software.

For somatic photostimulation, the center of the beam in the focal plane was directed at any point within the boundary of the soma as visualized under IR-DIC, but the precise three-dimensional overlap between the large irregularly shaped soma and the biconically convergent beam around the focal volume was not quantified. For dendritic and axonal photostimulation, two methods were used: the center of the beam was positioned over the structure by IR-DIC, using manual control of the servos, or a "side-scan" method in which flash photolysis of caged GABA was performed along a linear series of points spaced 1.5-2.0 µm apart, along an axis that was approximately orthogonal to the structure and that intersected it near the midpoint of the series. A least-squares fitting of the response amplitudes obtained by the side-scan provided a more accurate measurement of the response at the point of intersection. This method was used to confirm the accuracy of results obtained by IR-DIC positioning.

The effective beam diameter for photolysis in the focal plane was estimated by side-scan photostimulation of axons, taking advantage of their small diameter. The 2 width of a Gaussian fit to resulting plots of current amplitude versus position was 2.43 ± 0.36 µm at 1.24 ms (n = 19 scans from 2 cells). This width was similar to the diameter of the focused spot imaged by Lucifer yellow fluorescence (2.71 ± 0.05 µm, Gaussian fit to profile of CCD camera readout, with camera controller setting:  = 1).

Cell morphology

Live cell morphology was recorded on-line using a frame grabber board (PCI-1407, National Instruments) to capture a DIC image for each recorded response. Image-acquisition functions were integrated into the LabVIEW patch-clamp/motion-control program. Afterward, frames were cropped and tiled to reconstruct a z axis projection of the live cell, and an outline of the cell was traced and superimposed onto the photostimulation coordinates recorded by the XYZ positioning system. After each experiment, slices were fixed overnight at 4°C in phosphate-buffered saline with 2% glutaraldehyde, and the biocytin-filled cells were processed with the Vectastain Elite ABC kit and stained with a VIP peroxidase substrate kit (Vector Laboratories). Slices were cleared as whole-mounts in 80% glycerol, and fixed cell morphology was obtained by two methods: a Nikon Microphot microscope equipped with a CCD camera recorded images of sections of dendrites in different focal planes, which were cropped and tiled to yield a z-axis projection of the overall morphology; the XYZ positioning system and 63× objective were used to reconstruct the dendritic geometry in detail; images were captured by the frame grabber at a series of points along the dendrites separated by <10 µm, and the dendritic diameter at each point was estimated by a LabVIEW program that obtained mean densitometric profiles along parallel axes oriented orthogonal to the dendrite. The overall fixed-cell morphology was used to confirm that the recorded neuron was a mitral cell, to positively identify the dendrites as primary or secondary, and to verify that photolysis data were not complicated by dendritic branches of the recorded cell extending above and below the focal plane. The detailed reconstruction data were used in compartmental models of the dendrites to correct space-clamp errors (see following text).

Data analysis

Physiological responses were analyzed off-line with custom software written in LabVIEW and Origin 6.1 (OriginLab), and with the Mini Analysis Program (Synaptosoft). Corrections to pipette-bath liquid junction potentials were made using the JPCalc Program (Cell MicroControls).

To compare inward currents and spike modulation effects induced by different levels of caged GABA photolysis, data were correlated with a "flash-concentration" stimulus parameter: FC = (laser power at the cell) × (flash duration) × (caged GABA concentration) expressed in units of µJ.mM. The laser power at the cell is the nominal value calculated by multiplying unattenuated output power, as measured by the internal power meter, by the attenuation factors associated with the microscope optics. The FC value is a proportional measure of the total quantity of GABA released by photolysis in a fixed focal volume. It is proportional to peak concentration, provided that the rate of diffusion of photoproduct out of the focal volume, and local depletion of caged compound during the flash can be neglected.

Voltage-clamp data obtained by the side-scan method consisted of families of responses obtained from photolysis of caged GABA by 1.24-ms flashes, applied to a series of equally spaced points (0.5-1.5 µm apart) along an axis approximately orthogonal to a dendrite or axon. The current amplitude was measured at t_meas = 2 ms after shutter opening (i.e., 0.76 ms after shutter closing), around the middle of the rising phase of the response. The part of the rising phase after shutter closure and termination of photolysis represents the continued increase in current due to spatial summation of GABA-activated conductance as the photoreleased GABA spread locally by diffusion. Taking the larger current value at this time point reduced the relative contribution of whole cell noise to the error in measurement, at the expense of slightly reduced spatial resolution. Assuming an initial Gaussian distribution of photoreleased GABA of width w = 2 = 2.43 µm in the focal plane, and diffusion in two dimensions with coefficient D ~ 7 × 10⁶ cm²s¹, a time delay of t = 0.76 ms would reduce resolution by increasing the effective width to: w' ~ (4² + 8D.t) = 3.2 µm. In various high-resolution mapping experiments, consecutive side-scans were spaced on average 1.4-3.7 µm apart, a resolution similar to or finer than that set by diffusion. Each scan profile was fit to a Gaussian function by the Levenberg-Marquart algorithm to extract the peak amplitude. In a few cases, two local maxima were present in the scan profile. Examination of the biocytin reconstructed neurons showed that the second peak was correlated with a dendritic branch that was not visible under IR-DIC. In these instances, the second peak was removed manually from the data, and the remaining peak was fit.

Space-clamp errors associated with dendritic photolysis currents recorded under somatic voltage clamp were estimated by constructing compartmental models of recorded cells based on the dendritic geometry obtained from biocytin-stained cells. This error estimation was possible because the positions of dendritic photolysis sites are precisely known. Models of mitral cells were constructed in the NEURON simulator (Hines and Carnevale 1997). The soma was represented as a single cylindrical section (length, 20-30 µm; diameter, 20-25 µm), and the primary and secondary dendrites as segmented sections (compartment length, 1 µm) with piece-wise linear tapering. Taper intervals were variable, ranging from 15 µm proximally to 200 µm distally, depending on the cell and dendrite. In the primary dendrites, tapering was significant only proximally (less than ~40 µm from the soma), whereas in the secondary dendrites, tapering occurred along the length of the dendrite (Mori et al. 1983). For primary dendrites, mean diameter was 3.5 ± 0.6 µm at 20 µm from the soma and 3.0 ± 0.4 µm at 60 µm (n = 6 dendrites); for secondary dendrites, mean diameter was 2.7 ± 0.3 µm at 20 µm and 2.0 ± 0.3 µm at 60 µm (n = 8 dendrites). The model values of passive electrotonic parameters were taken as those determined in a recent modeling study to best-fit data from dual current-clamp recordings of action potentials in mitral cell primary dendrites (Shen et al. 1999): intracellular resistivity, R_i = 70  · cm; membrane resistance, R_m = 30,000  · cm²; membrane capacitance, C_m = 1.2 µF · cm². In the voltage-clamp experiments being simulated here, K⁺ was replaced by Cs⁺ in the intracellular solution, which would slightly reduce R_i by the ratio of the electrophoretic mobilities of the cations (~5%) (Hille 1984); this had a negligible impact on the computed space-clamp errors. To simulate the recordings of photolysis currents, a perfect voltage-clamp electrode was applied to the model soma, and the conductance activated by focal uncaging of GABA at a given position along a dendrite was simulated by locating an AlphaSynapse, g(t) = g_max · (t/) exp[  (t  )/], with zero reversal potential, on the corresponding dendritic compartment (at 1-µm accuracy). The value of  was set equal to the time-to-peak of the response to somatic photolysis (5-15 ms), and the somatic current was computed at the measurement time, t_meas < . This phenomenological model was able to closely fit the time course of the photolysis-activated conductance during its rising phase (when the current amplitude measurement was made), without explicitly modeling the diffusional spread of GABA at later times. The value of g_max that generated a measured somatic current was then obtained by iterative bisection of a conductance interval bracketing the measured current, and the predicted somatic voltage-clamp current with perfect space-clamp was calculated by multiplying the corresponding g(t_meas) by the holding potential.

Action potential amplitudes were measured as the difference between the voltages at two points: the spike peak determined by quadratic fit of five consecutive data points (50-kHz sampling) and a prespike inflection point, defined arbitrarily as the point where the second derivative of the voltage is equal to 1/10 of the local maximum value of the second derivative of the voltage on the rising phase of the spike (also located by 5-point quadratic fitting). This procedure always yielded a reproducible inflection point near the end of the depolarizing ramp preceding each spike. Spike width was measured as the time difference between half-peak amplitude points on the rising and falling phases of the action potential, the amplitudes being measured from the prespike inflection point. To reduce noise and improve the reliability of inflection point estimation, data arrays were smoothed with a second-order Chebyshev filter (ripple, 0.1 dB; cutoff frequency, 1.5 kHz). Applying this filter did not result in significant errors in the measurement of spike parameters: filtered action potentials with width 1.18 ± 0.03 ms (n = 9) had their peak voltages changed by a factor of 0.993 ± 0.002 relative to the unfiltered peaks, and their widths changed by a factor of 1.01 ± 0.02 relative to peaks filtered at 3 kHz.

For backpropagated dendritic action potentials, a backpropagation attenuation factor (BPAF) was defined for each spike, as the ratio of dendritically to somatically recorded amplitudes. The BPAF decayed after breakthrough into whole cell mode, and the initial value, BPAF_i, was estimated by extrapolation back to 1 min prior to breakthrough (the approximate time taken to achieve a dual recording by breakthrough at the soma, after initial breakthrough at the dendrite). Extrapolation was by linear or exponential fit to the monotonic change occurring during the first 1-10 min of recording. SDs were obtained from the 68% prediction interval for the least-squares fit. For each backpropagated spike, a backpropagation broadening factor, BPBF, was defined as the ratio of dendritically recorded to somatically recorded spike widths. The BPBF increased after breakthrough into whole cell mode and the initial value, BPBF_i, was also estimated by a similar back-extrapolation procedure. Backpropagation conduction velocity was computed from the time difference between the somatically and dendritically recorded action potential peaks and also decayed over time, so a similar extrapolation was applied to extract the initial conduction velocity. For each somato-dendritic pair of spike trains, the broadening and attenuation factors were calculated as the average BPAF and BPBF for the first three or four spikes (prior to the flash, if GABA was applied by photolysis).

When analyzing the effect of caged GABA photolysis on dendritic spike trains, flash timing usually fell between action potentials, so to correct for variations in spike timing relative to shutter opening, a corrected photolysis attenuation factor (PAF_d) for dendritic spike trains was estimated by back-extrapolating, to the flash time, the amplitude differences between the last preflash spike and a series of 4-10 postflash spikes. Referencing the postflash amplitudes relative to the last preflash spike was justified by the observed lack of activity-dependent attenuation in the dendritic spike train. Measurements were not taken from spikes that coincided with or overlapped a period of several milliseconds during or following the flash because during this period, both local GABA concentration and membrane voltage were changing rapidly. Back-extrapolation of amplitudes was performed by using the Levenberg-Marquart algorithm to fit an exponential decay to the postflash amplitude difference, and a standard error was estimated from the prediction band for 68% confidence level at the flash time. In cases where an exponential fit did not converge because the scatter in the postflash amplitudes masked the curvature in the plot of amplitude recovery versus time, extrapolation was performed by a linear fit, with standard errors in the extrapolated values taken from prediction bands at 68% confidence level. The effect of dendritic photoinhibition on somatic spike amplitude was too small to allow reliable curve fitting and extrapolation, and a somatic photolysis attenuation factor (PAF_s) was calculated directly by dividing the amplitude of the first postflash spike by the mean amplitude of all the preflash spikes. Again, this was justified by the observed lack of activity-dependent attenuation in the somatic spike trains. The standard error in PAF_s was obtained from the amplitude variance of the preflash spikes.

	RESULTS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Caged GABA photolysis activates a GABA_A receptor-mediated conductance in mitral cells

The membrane current evoked by photolytic release of GABA onto the mitral cell soma was recorded under whole cell voltage clamp while dialyzing with CsCl to block K⁺ conductances and shift the chloride reversal potential to near 0 mV. The bath included TTX and Cd²⁺ to block regenerative Na⁺ and Ca²⁺ currents. Under these conditions, laser flash photolysis of caged GABA evoked large transient currents that were inward at negative holding potentials (Fig. 1A). The amplitudes and decay rates of these currents varied considerably between cells: at 60 mV, with a 0.725-µJ flash (1.24 ms, FC = 0.29 µJ · mM), the peak inward current was 646 ± 267 pA (range, 275-1082 pA; n = 8 cells, 1 flash/cell), and decay time constants were 64 ± 30 ms (range, 36-119 ms, monoexponential fit to decay; n = 8 cells). Responses to laser flashes were not observed when caged GABA was omitted from the bath solution, and introducing 400-µM caged GABA into the bath did not activate a significant inward current in the absence of laser irradiation. The photolysis responses were quite stable during repeated stimulation of the same cell: the peak current recorded from one cell subjected to repeated somatic photolysis (n = 19 trials, 2 s apart) was 529 ± 11 pA, with the 2% variation being attributable to baseline noise in the whole cell current. The mean percentage variation in amplitudes for repeated stimulation of the soma and proximal dendrite was 3.9 ± 1.1% (average from n = 4 cells, 5 flash trials/cell). The current-voltage relation was nearly linear, with reversal potential near zero (E_rev = 1.9 ± 2.7 mV, n = 10 cells), consistent with activation of a chloride current (Fig. 1B). Responses were strongly blocked by 50 µM BMI (attenuation factor for peak current 0.057 ± 0.023, n = 4 cells at 60 mV; FC = 0.29 µJ · mM), indicating they were mediated by GABA_A receptors (Fig. 1C). The recovery kinetics were significantly slower at positive holding potentials (Fig. 1D): the decay time at +40 mV was 2.19 ± 0.55 times longer than the decay time at 40 mV (P < 0.001, paired t-test, n = 7 cells; monoexponential fit to decay). This slowing is consistent with the known voltage-dependent prolongation of decay kinetics of GABA_A receptor-mediated IPSCs (Otis and Mody 1992). The photolysis responses were most likely due to a direct action of the uncaged GABA on the membrane of the recorded mitral cells, without contributions from polysynaptic pathways involving glutamatergic excitation. Indeed, there were no significant differences, with or without 50 µM AP-5, 50 µM CNQX in the bath, between: decay times at 60 mV, reversal potentials, BMI attenuation factors, or the ratio of recovery kinetics at ±40 mV (P > 0.1, n = 5).

View larger version (15K): [in this window] [in a new window]   Fig. 1. Mitral cell response to GABA photostimulation. A: membrane currents recorded from a mitral cell under whole cell voltage clamp in response to somatic flash photolysis of 400-µM O-CNB-caged GABA (3 ms, 0.88 µJ). Nominal holding potential was varied from 80 to +40 mV in 10-mV steps. Dialysis of the cell with a CsCl-based pipette solution reversed the chloride currents carried by the GABA receptor at negative membrane potentials. The bath contained 1 µM TTX, 100 µM Cd2+. B: current-voltage relation for the data in A. Current amplitudes were measured 8 ms after shutter opening, before the peak of the current. The nominal voltage was corrected for the calculated pipette-bath liquid junction potential (5.6 mV). C: block of the inward current in A at 70 mV by bath perfusion of 50 µM bicuculline methiodide (BMI). D: exponential fits to the overall decay of photolysis responses in (A) at 40 mV and +40 mV (fit curves have time constants 47 and 96 ms, respectively).

Spatial distribution of currents activated by caged GABA photolysis

Flash photolysis was applied at different points on the mitral cell membrane to determine the spatial distribution of the receptors underlying the GABA-activated inward current recorded somatically under whole cell voltage clamp with CsCl dialysis. Figure 2 shows typical data obtained from a mitral cell (photolysis sites 1-3 µm apart), in which the primary dendrite was mapped out to 170 µm from the soma, and the secondary dendrite out to 140 µm (the maximum distances at which a current was detectable by somatic recording). For both dendrites, the response was largest proximally and declined progressively with increasing distance from the soma. Exponential fits to these declines yielded decay lengths of 75 ± 28 µm for primary dendrites (range, 24-95 µm; n = 6 cells) and 78 ± 48 µm for secondary dendrites (range, 42-166 µm; n = 7 cells). The responses evoked by dendritic stimulation were comparable in time course to somatic responses (Fig. 2B, 1 and 2) and were also blocked by BMI (data not shown).

View larger version (61K): [in this window] [in a new window]   Fig. 2. Mapping of GABA receptors on mitral cell dendrites. A, top: the x-y coordinates of photostimulation points along the dendrites of a mitral cell (), 41 points on the secondary dendrite (left) and 58 on the primary dendrite (right), overlayed onto a tracing of the Nomarski image of the cell (position of the somatic recording electrode is indicated). Scale bar: 50 µm. Bottom: plots of the magnitudes of the inward currents evoked by local photolysis of 270-µM O-CNB-caged GABA (3.5-ms flashes, 0.4 µJ), measured at the end of shutter opening (); also shown are the same currents after compensation for space-clamp errors (). Superposed over the same abscissas are plots of the percentage space-clamp error (% SCE: ) and the space-clamp error-compensated currents divided by the dendritic diameter that provide a measure of the local conductance density (). The cell was dialyzed for 90 min with a CsCl-based pipette solution, and the nominal holding potential was 60 mV. The abscissas in these plots represent integrated distance along the dendrites, (dx2 + dy2 + dz2), with 0 distance taken at a point where the base of the dendrite joins the soma. B: raw data showing photostimulation responses evoked at each of the points plotted in A. Numbers on the left of every 10th trace are also labeled on the corresponding photolysis sites and amplitudes in A. B, 1 and 2: responses along the secondary (1) and primary (2) dendrites superimposed by rescaling to show similarity in time courses. Integrated distances for traces in B1 are (in µm): 0, 23.4, 46.5, 64.4, 90.8, 101.1, 109.5, 118.2, and 130.8; for traces in B2 are (in µm), 0, 29.1, 54.6, 76.7, 105.6, and 161.2. C: bright-field image of the whole-mount of the fixed slice containing the cell in A and B visualized by biocytin staining. The approximate locations of the distalmost photostimulation points, 41 on the secondary dendrite and 58 on the primary dendrite, are marked. Scale bar: 100 µm.

One factor that might contribute to these declines is a reduced ability of the somatic electrode to clamp the dendritic membrane voltage at more distal locations (Bhalla and Bower 1993; Spruston et al. 1993). The contribution of space-clamp errors to the spatial profile was estimated by numerical simulation of reconstructed cells and was found to be relatively small ( and  in plots of Figs. 2A, 3A, and 4, A and B). Estimated space-clamp errors increased approximately linearly with increasing distance from the soma (, Fig. 2A). For primary dendrites, the relative error was 2.7 ± 1.7% at 20 µm from the soma and 6.8 ± 4.2% at 60 µm (n = 6 dendrites); for secondary dendrites, it was 2.5 ± 0.9% at 20 µm and 7.5 ± 2.9% at 60 µm (n = 8). The computed space-clamp errors were modest because the photolysis locations were <200 µm from the soma, where both dendrites are fairly wide (diameter of primary dendrite more than ~3 µm, of proximal secondary dendrite more than ~1.5 µm), the assumed intracellular resistivity was relatively low (70  · cm), and the rising phases of the photolysis-activated currents were relatively slow (times to peak, ~5-15 ms). The similar time courses of the responses along the dendrites (Fig. 2B, 1 and 2) is consistent with a weak effect of cable filtering. After compensation for space-clamp errors, the mean exponential decay lengths were increased by 12% (to 84.3 ± 31.4 µm) for primary dendrites (n = 6) and 28% (to 100 ± 64 µm) for secondary dendrites (n = 8).

The spatial profiles of the space-clamp compensated responses were divided by the dendritic diameters to obtain a proportional measure of the GABA-activated conductance per unit membrane area (Figs. 2A, 3A, and 4, A, B: plotted as ). This revealed a significant difference between the conductance density profiles of the primary and secondary dendrites. In primary dendrites, dividing by the diameter made the profile more shallow proximally, where the dendrite tapers off from the soma (less than ~30 µm) but had much less effect on the profile at more distal locations where the diameter is more uniform. Decay lengths for exponential fits to primary dendritic profiles at >30 µm from the soma were increased by only 16 ± 14% (means increased from 80 ± 48 to 91 ± 49 µm, n = 6) after diameter division. Linear regression indicated that the conductance density of the primary dendrite decreased significantly with increasing distance from the soma (slope, 11.3 ± 9.3 pS · µm²; P < 0.01, n = 6 dendrites, probed lengths 50-300 µm). By contrast, in secondary dendrites, which taper continuously, dividing by the diameter strongly reduced the decay rate of the profile of conductance versus distance. For half of the dendrites tested, exponential decay lengths were increased by 43 ± 10% (means increased from 94 ± 64 to 131 ± 81 µm, n = 4), and linear regression of the conductance density profile revealed a significant residual decay (slope, 10.3 ± 7.9 pS · µm²; P < 0.05, n = 4 dendrites, probed lengths 50-150 µm); in the remaining four dendrites, the density profiles were too shallow to allow exponential fitting, and linear regression did not indicate a significant decrement of conductance density with distance (1.6 ± 5.6 pS · µm², P > 0.05, n = 4 dendrites; probed lengths, 60-90 µm).

The recorded currents exhibited variations in amplitude as a function of distance along the more proximal parts of the dendrites (Fig. 2A), which contributed to the scatter in decay lengths obtained from exponential fits to the overall decay. A possible source of such variation is random error in IR-DIC-guided positioning of the laser focus onto the dendrite. To reduce the contribution of random positioning errors, data were also acquired by the side-scan method, and a least-squares fit of the scan profiles was used to estimate the maximum amplitude on the dendrite. The accuracy of the method was assessed by repeating side scans across a fixed location on a dendrite (Fig. 3C). The average error in peak amplitudes obtained from Gaussian fits to four repeated scans was 10 ± 7% (n = 3 cells; 2 primary and 1 secondary dendrite; peak current range, 64-144 pA). Raw data from sequential side-scans of a proximal section of secondary dendrite are shown in Fig. 3B, and the corresponding peak amplitude estimates obtained by fitting are plotted in Fig. 3A. These results show that the somatically recorded current activated by local GABA photostimulation along the dendrite (with 3.2-µm resolution) is nonuniform with local peaks and valleys. These local peaks were still present after the data were corrected for space-clamp errors, and the dendritic taper had been taken into account by dividing by the diameter (Figs. 3A and 4, A and B). This indicates that there are sites along the dendrites where the density of the GABA-activated conductance is locally elevated.

View larger version (39K): [in this window] [in a new window]   Fig. 3. Spatial dependence of GABA-activated currents revealed by side-scan photolysis. Mitral cells were recorded under somatic whole cell voltage-clamp (60 mV) with CsCl dialysis, and the bath artificial cerebrospinal fluid (ACSF) contained 2 µM TTX, 200 µM Cd2+, 50 µM AP-5, and 50 µM CNQX. Serial photolysis of 400-µM O-CNB caged GABA (1.24-ms flash duration, 0.73 µJ) was performed by moving the center of the focal volume along linear tracks approximately orthogonal to the secondary dendrite. A: IR-DIC tracing, projected onto x-y plane, of a recorded mitral cell showing primary dendrite (PD) and secondary dendrite (SD). The recording pipette is shown at the soma, and the series of  overlayed along the secondary dendrite indicates the x-y coordinates of peaks obtained from Gaussian fits to side-scan photolysis response profiles (scale bar: 20 µm). The plot above these points shows the corresponding peak amplitudes as a function of position, before () and after () compensation for space-clamp errors. The un-normalized conductance density (compensated current divided by diameter) is plotted on the same abscissa (). *, putative local peaks in the amplitude profile. The left-most peak satisfied the spatial average nonmonotonicity criterion (P < 0.05). Total integrated distance along the dendrite covered by 36 side-scans was 73.7 µm, and mean interscan distance was 2.0 µm. The plot abscissa represents integrated distance (dx2 + dy2 + dz2), which is larger than the x-y projection distance. B: raw side-scan data from the cell in A. Magnitude of inward current at 2 ms after shutter opening is shown as line plots of y-axis scans, stacked along the x axis. The secondary dendrite is approximately parallel to the x-axis. C: raw data and Gaussian fits to 4 side-scans repeated across the 1 location on a secondary dendrite. The scatter in the peak amplitudes of these fitted curves is 4.7%. The relative error for side scans of dendrites from n = 3 cells was 10 ± 7%. The error bars on the plots reflect this error estimate.

A nonmonotonicity criterion based on spatial averaging was used as a simple test for local peaks. One or several consecutive photolysis points along a dendrite was deemed a local peak if there was a series of consecutive points closer to the soma (the "valley" points) whose mean (spatial average) amplitude was significantly less than the amplitude (or mean amplitude) of the test point(s) (P < 0.05; t-test, independent samples). This test was applied to both side-scan data, and IR-DIC data assuming random targeting errors were independent of position. For the side-scan data, it is expected to underestimate the number of local peaks because the spatial variance was larger than the estimated 10% error in Gaussian fits. Applying this test to secondary dendrites (Fig. 4B), 10 local peaks were detected (5 with P < 0.05, 3 with P < 0.01, 2 with P < 0.0005; mean valley/peak amplitude ratio, 0.57 ± 0.13), from a total of seven cells (6 side-scan, 1 IR-DIC; maximum distances 53-147 µm from soma; summated length over all cells 579 µm); on primary dendrites (Fig. 4A), 9 local peaks were detected (5 with P < 0.05, 4 with P < 0.005; mean valley/peak amplitude ratio, 0.62 ± 0.07), from a total of four cells (3 side-scan, 1 IR-DIC; maximum distances 54-300 µm from soma; summated dendritic length over all cells 830 µm).

View larger version (28K): [in this window] [in a new window]   Fig. 4. Spatial dependence of GABA-activated currents along mitral cell axons and dendrites. Currents were recorded from mitral cell somata by whole cell voltage-clamp with CsCl dialysis, and primary (A) and secondary (B) dendrites, and axons (B and C) were probed by side-scan photolysis of 400-µM O-CNB caged GABA, as described in Fig. 3. Tracings of the cells from IR-DIC images, projected onto the x-y plane, are shown, with  positioned at the sites of side-scan peaks (for clarity, a circle is drawn only at every 2nd site). Scale bars: 50 µm. , the beginning and end of the dendritic or axonal segment probed by photolysis. Total integrated distances and mean interscan distances were (in µm): A, 124.6, 2.3; B, secondary dendrite: 58.5, 2.0; axon: 34, 1.9; C, 31.0, 1.48. The plots display the peak current amplitudes as a function of distance from the soma, for the raw data (), and after compensation for space-clamp errors (). The un-normalized conductance density (space-clamp compensated current divided by diameter) is also plotted on the same abscissa (). *, putative local peaks in the amplitude profile satisfying the spatial average nonmonotonicity criterion (P < 0.05). PD, primary dendrite; SD, secondary dendrite; Ax, axon.

Sometimes it was possible to visualize and trace the axons of the mitral cells under IR-DIC illumination, and a few of these were also probed for GABA responses. Figure 4, B and C, illustrates side-scan data obtained from axons, showing local peaks and valleys in the profiles. A total of four local peaks were detected on axons (1 with P < 0.05, 3 with P < 0.01; mean valley/peak amplitude ratio, 0.54 ± 0.03), from a total of three cells (all by side-scan; maximum distances 19-34 µm from soma; summated length from all cells 84 µm). Compared to dendrites, the amplitudes of currents evoked by axonal photostimulation declined on average more strongly with increasing distance from the soma. The mean decay length from exponential fits to axon profiles from n = 3 cells was 29 ± 4 µm (range, 25-31 µm). This mean includes the profile shown in Fig. 4B but not that in Fig. 4C, which was not well fit by an exponential decay due to the large spatial fluctuations. Space-clamp corrections are not shown for the axon data because of the difficulty in accurately quantifying the small diameter of the axon initial segment, and the uncertainty in the R_m value. Trial calculations showed that predicted space-clamp errors were quite sensitive to diameter variations around ~1-2 µm and to the unknown distribution of a large leak conductance (R_m = 1,000  · cm²) along the initial segment (Shen et al. 1999).

Currents activated by caged GABA photolysis are comparable in magnitude to IPSCs from presynaptic spines

To address the question of whether caged GABA photolysis induces an unphysiologically large-conductance change, currents evoked by photolysis at the soma were compared with currents induced by GABA released from presynaptic spines of dendrodendritic synaptic contacts at the same location. Making the comparison at the soma rules out any cable filtering of dendritic postsynaptic currents caused by loss of voltage clamp. Spine activation was spatially restricted by local photolysis of caged glutamate at the soma, and the bath ACSF contained 0 added Mg²⁺ to facilitate activation of N-methyl-D-aspartate (NMDA) receptors. Figure 5A shows a series of currents recorded from a mitral cell dialyzed with CsCl with 1 µM TTX in the bath. Baseline control recordings (top) exhibited a low rate of spontaneous postsynaptic events. These events could be blocked by 50 µM BMI, indicating that they were reversed IPSCs mediated by GABA_A receptors (Kirillova and Lin 1998; Wellis and Kauer 1993). Glutamate photostimulation evoked a prolonged barrage of asynchronous events, continuing >1 s. This barrage became more intense as more glutamate was released by higher laser energies, and it was abolished by 50 µM BMI (bottom). This indicated that the evoked events were reversed IPSCs mediated by GABA_A receptors and that photostimulation triggers local GABA exocytosis from presynaptic spines. After addition of BMI, a residual slow inward current remained that was blockable by 60 µM AP-5 (data not shown), indicating involvement of NMDA autoreceptors (Petralia et al. 1994). The caged glutamate responses resembled other responses caused by asynchronous release of GABA onto the mitral cell: i.e., responses evoked by puffer pipette application of NMDA or KCl depolarization of the inhibitory interneurons (Friedman and Strowbridge 2000), stimulation of the mitral cell by voltage pulse (Isaacson and Strowbridge 1998; Schoppa et al. 1998; Wellis and Kauer 1993) or uncaging calcium in the mitral cell to release glutamate (Chen et al. 2000; Isaacson 2001). Figure 4B shows the results of an analysis of the glutamate-evoked IPSCs recorded from the cell in Fig. 4A. Mean event amplitude was 103.5 ± 81.6 pA, with many events in the 200- to 300-pA range; mean decay time was 13.6 ± 10.6 ms. Analysis of data from a second cell yielded larger amplitudes, and similar decay kinetics (Fig. 5B, right).

View larger version (31K): [in this window] [in a new window]   Fig. 5. Comparison of GABA photostimulation-evoked currents and inhibitory postsynaptic currents (IPSCs) evoked by spine activation. A: GABAergic IPSCs evoked by spine activation, recorded from the mitral cell soma under whole cell voltage-clamp at 60 mV, with CsCl dialysis. The bath included 1 µM TTX to block regenerative Na+ currents. Top: a control recording with no photostimulation. Lower traces: responses to photolysis of 320 µM -CNB-caged glutamate at a site on the soma (5 ms flashes, 0.37-2.25 µJ). Bottom: the response to caged glutamate photolysis (5 ms, 2.25 µJ) with 50 µM BMI in the bath. B, left: event histograms of IPSC amplitude (bottom, 40-pA bins) and IPSC decay time (top, 5-ms bins) for the first 1-s period of glutamate responses (5-ms flash, 2.25 µJ) recorded from the cell in A: (cell 1, mean amplitude, 103.5 ± 81.6 pA; mean decay time, 13.6 ± 10.6 ms; n = 57 events). Right: similar event data recorded from another cell (cell 2, n = 114 events) with a more sustained response (2.4 s; 3 repeats; stimulus, 7 ms, 2.0 µJ). The number of events at larger amplitudes was greater (mean, 171.1 ± 122.4 pA), and the mean decay time was similar (9.8 ± 6.9 ms). C: comparison of IPSCs evoked by glutamate photostimulation (top, same as 2.25 µJ trace in A), and inward currents evoked by GABA photostimulation (270 µM, 5 ms, 0.22-2.2 µJ) at the same location on the soma of the mitral cell in A. The bath included 1 µM TTX. D: plot of the magnitude of the inward current at 4 ms after shutter opening, as a function of flash energy, for the family of GABA photostimulation-evoked currents in C. The data were fit to a Hill equation: inward current, I = Imax · FCn/{kn + FCn}, where FC = (flash energy) × (caged GABA concentration). Fit parameters were: k = 0.37 ± 0.02 µJ · mM, Imax = 562 ± 26 pA, n = 1.7 ± 0.1.

After recording glutamate responses, the bath solution was switched to ACSF containing TTX and caged GABA, and GABA was uncaged at the same somatic site. Figure 4C compares the postsynaptic events recorded from the cell of Fig. 4A with GABA-activated currents recorded from the same cell over a range of flash energies. Uncaging of GABA resulted in currents that were several hundred picroampere in amplitude; the dose-response relationship could be fit to a Hill equation: I = I_max · FCⁿ/{K_1/2ⁿ + FCⁿ}, K_1/2 = 0.37 ± 0.02 µJ · mM, I_max = 562 ± 26 pA, n = 1.7 ± 0.1. Hill fits to dose-response relationships for the somatic GABA-activated current obtained from three additional cells gave consistent values for parameters K_1/2 (0.36 ± 0.04 µJ · mM; range, 0.31-0.40 µJ · mM, n = 4) and n (1.85 ± 0.14; range, 1.69-2.03, n = 4), which characterize the GABA receptor, whereas I_max varied widely (522-1,436 pA). The wide variation in maximal somatic current probably reflects in part the technical difficulty in specifying the three-dimensional geometry of overlap between the photolysis beam and the irregularly shaped somatic membrane.

The peak GABA concentration associated with FC = K_1/2 ~ 0.37 µJ · mM is difficult to know precisely because the EC₅₀ of GABA_A receptors under nonequilibrium conditions (as might occur during flash photolysis) depends on entry of receptors into desensitized states. For the currents measured several milliseconds after photolysis, the GABA concentration corresponding to FC = K_1/2 might lie between EC₅₀ ~ 10-40 µM at equilibrium (Feigenspan et al. 2000), and EC₅₀ ~ 185 µM for 1-ms transient pulses (Galarreta and Hestrin 1997). If the receptors are assumed to be near equilibrium, a rough estimate of the peak GABA concentration after uncaging can be deduced from the observed attenuation of the peak current by the competitive inhibitor bicuculline. Assuming a Hill coefficient of 1 for the GABA_A receptor (Feigenspan et al. 2000; Ueno et al. 1997), at a bicuculline concentration of 50 µM, an attenuation factor of ~0.06 corresponds to IC₅₀ ~3.2 µM, which lies between the IC₅₀ values at 10 µM GABA (1.6 µM) and 30 µM GABA (5.8 µM) for the GABA_A receptor with subunit composition 122 (Ueno et al. 1997). The latter data are applicable here because mitral cells strongly express mRNA for 1, 1, 2, 3, and 2 subunits of the GABA_A receptor (Laurie et al. 1992), and receptors with different subunits have similar affinities for GABA and bicuculline (Ebert et al. 1997; Krishek et al. 1996). Interpolation between the IC₅₀ values gives ~20 µM GABA as a peak concentration. This number is consistent with the cited range of EC₅₀ values at equilibrium because the attenuation by BMI was measured at FC = 0.29 µJ · mM, only slightly below the K_1/2 of the photolysis responses. Thus an equilibrium approximation may be useful for describing the rising phases of photolysis responses, which are relatively slow (5-15 ms) compared with unitary IPSCs from granule cells (rise times, <1 ms).

The tests with caged glutamate showed that the magnitudes of the inhibitory conductances resulting from the uncaging of GABA can be comparable to the range of inhibitory conductances of IPSCs received from presynaptic spines. Photolysis with FC values of ~0.45-0.90 µJ · mM generated inward currents of ~300-600 pA, which is in the range of the amplitudes of individual IPSC events evoked by direct activation of NMDA receptors on spines. A significant difference was the slower decay of GABA photostimulation currents compared with spine-evoked IPSCs (Fig. 4C). Spine IPSCs are expected to decay more rapidly because the GABA released into the synaptic cleft is much more localized than the GABA released by photolysis. The peak concentration of GABA in the synaptic cleft following exocytosis may be more than ~500 µM (Jones and Westbrook 1995; Maconochie et al. 1994). This is considerably higher than would be attainable by flash photolysis of 400-µM caged GABA, which may generate peak transients of only ~10-100 µM. However, the uncaging of GABA can activate conductances of comparable magnitude by spatial summation over a larger membrane area.

The slower time course of the conductance change produced by uncaging GABA might actually be a better simulation of physiological conditions. Coordinated firing of granule cells in vivo during oscillations of populations of mitral and granule cells (Freeman and Baird 1987; Li and Hopfield 1989) would be expected to result in strong spine depolarization by backpropagating action potentials in the granule cell apical dendrites. This would provoke stronger, more synchronous GABA release driven by voltage-sensitive calcium channels, facilitating a temporal summation of IPSCs and magnifying and prolonging the inhibitory input to mitral cell dendrites. Focal depolarization of spines by KCl has been shown to activate a large (>1 nA), slow (>100 ms) Cd²⁺-sensitive dendrodendritic IPSC in mitral cells (Halabisky et al. 2000), similar to IPSCs evoked by GABA photostimulation at higher laser power. IPSCs much larger than 1 nA can also be evoked by glomerular shock (Schoppa et al. 1998). Under such conditions, it is expected that IPSCs received from granule cells and other interneurons would be more closely mimicked by the currents evoked here by uncaging of GABA.

Inhibition of somatic action potentials by photolysis of caged GABA

The mitral cell soma initiates spike trains in response to depolarizing current conducted from the primary dendrite during synaptic excitation of the apical dendritic tuft. The EPSC from the primary dendrite is counteracted by GABAergic inputs on the soma and secondary dendrites, activated during self- and lateral inhibition. In the voltage-clamp recordings described in the preceding text, functional GABA receptors were found distributed over the secondary dendrites 150 µm from the soma. At what range do these receptors influence action potential firing at the soma? In general, it is expected that distal inhibition would be less effective at shunting the somatic EPSC, but the actual range depends on the density of the GABA receptors and the magnitude of depolarizing current. This was demonstrated by recording from mitral cells under somatic current clamp, dialyzing with low internal chloride solution, and initiating spike trains by injecting square current pulses into the soma. During the spike trains, inhibitory input was applied locally along the dendrite by caged GABA photolysis, using IR-DIC imaging to position the laser focus on the dendrite (Fig. 6B).

View larger version (40K): [in this window] [in a new window]   Fig. 6. Range of somatic inhibition in the mitral cell secondary dendrite. A: families of responses recorded from the soma of a mitral cell under whole cell current clamp (52 mV resting potential), while injecting 200-ms current pulses of different magnitudes (left to right column: 50, 100, and 150 pA) and applying GABA photostimulation (270-µM caged GABA, 7.2 ms, 3.2 µJ) at different points along the secondary dendrite. On the left, stimulus points are numbered, and the corresponding integrated distances from the soma [(dx2 + dy2 + dz2)] are indicated in µm. Bottom traces are controls without photostimulation. , timing of photolysis. B: the x-y coordinates of photostimulation points along the secondary dendrite of the cell in A overlayed onto a tracing of the Nomarski image of the cell (position of the recording electrode is indicated). The dendrite extending to the right is the primary dendrite, which could be followed up to the glomerular layer with IR-DIC imaging. Scale bar: 50 µm. C: collapse of the range of somatic inhibition with increasing strength of excitation. The histogram plots the maximum observed distance at which GABA photostimulation was able to terminate a 200-ms spike train evked by injected current pulses of various amplitudes.

Figure 6A illustrates the different spatial patterns of inhibition obtained when varying the injected current at a fixed level of photolysis (FC = 0.85 µJ · mM). For 50-pA current pulses, barely suprathreshold for repetitive firing (40 pA was subthreshold), the spike train was terminated by focal inhibition at points extending out as far as 137 µm from the soma (left); at 100 pA (middle), inhibition applied further than 78 µm from the soma failed to terminate somatic firing; and at 150 pA (right), only inhibition on the most proximal site (8 µm) was effective in terminating the spike train. A larger depolarizing current requires a larger conductance shunt to prevent spiking. Thus the decrease in the maximal spatial range of spike termination with higher somatic current (Fig. 4C) shows that distal inhibition is indeed less effective at shunting somatic current. For the purposes of comparing data from different cells, the range of spike termination was defined arbitrarily as the maximal range for termination of repetitive firing without recovery of spiking during the 100-ms time period after GABA release. Trains of action potentials exhibited spike frequency adaptation, so the release of GABA was timed at 100 ms after the beginning of the current pulse, when frequency adaptation was largely complete (mean adaptation time constant, 49 ± 26 ms, exponential fit, n = 6 cells). The spike frequency adaptation was present when CNQX and AP-5 were included in the bath to block self inhibition by dendrodendritic feedback.

Different cells exhibited different firing thresholds depending on their input impedance and resting potential; so to compare data from several cells, the injected current was corrected by subtracting out a threshold current estimated from subthreshold pulses applied to each cell. Data from several cells confirmed that at approximately equal levels of photolysis, the maximal range of spike termination was negatively correlated with threshold-corrected current (r = 0.57, range slope = 0.33 ± 0.11 µm/pA¹, n = 4 cells; FC = 0.75-0.91 µJ · mM). Conversely, at approximately equal magnitudes of current, the maximal range of spike termination was positively correlated with the level of photolysis (r = 0.70, range slope = 1.8 ± 0.8 µm. µJ¹ · mM¹, n = 5 cells; I_corrected = 90-110 pA). The maximum observed range (<140 µm) was limited by the technical difficulty of visualizing the finely tapered secondary dendrites at more distal locations. The considerable cell-to-cell variability in these range measurements may be caused by several factors, such as the spatial heterogeneity in GABA receptor density along the secondary dendrite (Figs. 3 and 4) and the high sensitivity to shunting when the somatic membrane potential lies close to firing threshold. Spatial heterogeneity in dendritic conductance would also contribute to the patchiness of the spatial profiles of inhibition shown in Fig. 6.

Action potential backpropagation in the secondary dendrites

Beyond a certain range, which depends on the relative levels of excitation and inhibition, distal GABAergic inhibitory inputs to the secondary dendrite are ineffective at blocking somatic action potentials. However, GABAergic granule-mitral synapses are paired with reciprocal glutamatergic mitral-granule synapses through which the distal dendrite can send output to granule cells. Activation of these outputs requires sufficient distal depolarization to activate presynaptic Ca²⁺ channels. Such depolarization could be provided by laterally backpropagating action potentials. In the mitral cell primary dendrite, apically backpropagating action potentials are well characterized because of the ease of recording from the relatively thick dendritic trunk (Bischofberger and Jonas 1997; Chen et al. 1997; Shen et al. 1999). The finer, tapered secondary dendrites have only recently been shown to support active backpropagation of action potentials (Charpak et al. 2001; Margrie et al. 2001).

Dual whole cell current-clamp recordings from soma and dendrite confirmed that trains of action potentials initiated at the soma do backpropagate laterally into the secondary dendrite (Fig. 7). In these recordings, dendrodendritic feedback inhibition was blocked by glutamate receptor antagonists. The backpropagated spikes in the secondary dendrite were found to be attenuated relative to the somatic spikes. The backpropagation attenuation factor (BPAF) decayed monotonically after breakthrough into whole-cell mode at the dendrite (decay rate, 0.04 ± 0.02 min¹; range, 0.02-0.07 min¹; linear regression, n = 5 cells). This decay was usually due to a faster decay in the amplitude of the dendritically recorded spike, relative to the more stable somatically recorded spike (somatic, 0.19-0.48 mV/min; dendritic, 0.09-4.75 mV/min; n = 4 cells). In one other cell, the BPAF decay was also due to a slow growth in the somatic spike (0.67 ± 0.10 mV/min). The initial attenuation factor (BPAF_i), estimated by back-extrapolation (see METHODS) was 0.75 ± 0.07 (range 0.70-0.86; P < 0.005, t-test for BPAF =1; n = 6 cells) at distances of 93-152 µm (mean 112.3 µm) from soma. The backpropagated spikes were slightly broadened, and the backpropagation broadening factor (BPBF) increased following breakthrough (rate 0.06 ± 0.04 min¹, range 0.006-0.1 min¹, linear regression, n = 5 cells). The estimated initial broadening factor (BPBF_i), was 1.19 ± 0.10 (range, 1.06-1.29; P < 0.05; n = 6 cells). For each cell, the gradual broadening of dendritic spikes after whole cell breakthrough was directly correlated with attenuation: linear regression of BPBF against 1-BPAF gave a mean correlation coefficient of r = 0.962 ± 0.039 (BPAF range, 0.4-0.86, n = 5 cells). Like the amplitudes, the gradual increase in relative broadening was due primarily to an increase in the width of dendritically recorded spikes with the somatic spike widths being more stable. The peak of a dendritically recorded spike always occurred after the peak of the corresponding somatically recorded spike, showing that action potentials were initiated at the soma. Backpropagation conduction velocities calculated from the peak time difference exhibited a gradual decrease during whole cell recording, and the extrapolated initial velocity (BPCV_i) was 0.44 ± 0.14 m/s (n = 6 cells).

View larger version (18K): [in this window] [in a new window]   Fig. 7. Action potential backpropagation in the mitral cell secondary dendrite. A: whole cell current-clamp recording of a train of action potentials recorded simultaneously from the soma (left), and the secondary dendrite (right) at a point 152 µm from the soma. Action potentials were evoked by a 200-ms, 200-pA current pulse injected into the soma. Mean firing frequency is 39.3 Hz (initial, 50 Hz; final, 38.6 Hz). The pipettes contained K-methylsulfate, and the bath solution contained 60 µM 60 µM 2-amino-5-phosphonopentanoic acid (AP-5) and 10 µM 6-cyano-7-nitroquinoxalene-2,3-dione (CNQX) to block dendrodendritic feedback inhibition. Resting potential was 48 mV. In this cell, intrinsic membrane potential oscillations were visible. B: the x-y projection of the recorded cell, reconstructed by biocytin staining. The positions of the somatic and dendritic recording sites are shown are shown by pipette symbols. Scale bar: 100 µm. C: expanded plots of the first (left) and last (right) action potentials in A, with somatic and dendritic recordings superimposed. D: plot of initial backpropagation attenuation factor (BPAF) vs. distance from the soma (data are from dual whole cell recordings from the soma and secondary dendrite of n = 6 mitral cells).

Trains of backpropagating action potentials in the secondary dendrite did not exhibit the dramatic activity-dependent attenuation that has been reported for spi