At P0, the frontal cerebral cortices of both hemispheres from one pup per litter were used for expression analysis (n = 3–5 different litters). The dorsal cortex was dissected according to Khazipov et al. (2015), ∼1.8 to 0.4 mm from bregma (Figure 1A).

The frontal cortex was placed in 1 ml of TRIZOL^® reagent (Life Technologies, Bartlesville, OK, United States) and stored at −70°C until required. To obtain total RNA, the tissue was homogenized by pipetting, incubated for 15 min at 30°C, chloroform was then added, and samples were vigorously shaken for 15 s and then incubated for 3 min at room temperature. After centrifugation (1,700 × g) for 15 min at 4°C, the chloroform phase was transferred to a new tube, isopropanol was added, the tubes were vigorously shaken, and samples were incubated at 30°C for 10 min before centrifugation (11,000 × g) at 4°C. The supernatant was then removed, and the pellet was washed twice for 5 min with 1 ml of ethanol.

Total RNA was resuspended in 20 μl of RNAse-free water and incubated for 10 min at 60°C; RNA integrity was determined by the presence of 18S, 28S, and 5S ribosomal RNAs, after the electrophoresis of 1 μl of total RNA in 2% agarose gels.

For reverse transcription (RT) reactions, 1 μg of total RNA was mixed with 0.5 μg of random hexamers, 0.5 μl of a mixture of dinucleotide triphosphates (dNTPs, 10 mM), 2 μl of reverse transcription buffer (10×), 0.5 μl of ribonuclease inhibitor (40 U/μl), 4 μl of MgCl₂ (25 mM) and 5 U of AMV Retrotranscriptase (Promega, Madison, WI, United States) in a total volume of 20 μl with RNAase-free water. The mixture was incubated for 60 min at 42°C, the reaction was stopped for 5 min at 95°C, and the cDNA was kept at −20°C until required.

End-point PCR was performed to verify the size of the products. Dynamic ranges were performed for each transcript using the KAPA™ master mix Syber Fast qPCR^® (KAPA, Bartlesville, OK, United States), 1 μl of pre-amplified cDNA from 1, 10, 100, and 1,000 ng of cDNA to determine detection threshold, efficacy, slope, and to ensure a single fragment of amplification melting curves were obtained to observe a single peak (Kubista et al., 2006).

For qPCR, 1 μl of 500 ng of pre-amplified cDNA was assayed to obtain the relative expression of Satb2, Tbr1, and FoxP2, with GAPDH (glyceraldehyde 3-phosphate dehydrogenase) as an endogenous control. The sequences of the primers employed were those previously reported by Valle-Bautista et al. (2020):

Satb2 Forward (F): 5′-CCGCACACAGGGATTATTGTC-3′; reverse (R): 5′-TCCACTTCAGGCAGGTTGAG-3′, Tbr1 F: 5′-GGAAGTGAATGAGGACGGCA-3′; R: 5′-TGGCGTAGTT GCTCACGAAT-3′, FoxP2 F: 5′-GAAAGCGCGAGACACATCG GAPDH-3′; R: 5′-GAAGCCCCCGAACAACACA-3′, and GAPD H F: 5′-GGACCTCATGGCCTACATGG-3′; R: 5′-CCCCTCCT GTTGTTATGGGG-3′.

Reactions were performed with an initial denaturation period of 2 min at 95°C, followed by 40 denaturation cycles of 30 s at 95°C, alignment for 15 s at 63°C (FoxP2), 60°C (Tbr1), 57°C (Satb2), or 58°C (GAPDH). At the end of the incubation, melting curves were obtained. Results were analyzed using the 2^–ΔΔCT relative expression method (Livak and Schmittgen, 2001).

One or two animals (P0 and P21) per litter were used (n = 3–6 different litters). P0 and P21 rats received 50 mg/kg of pentobarbital intraperitoneally (Laboratorios PiSA) and were transcardiacally perfused with phosphate-buffered saline (PBS), pH 7.4, followed by Boüin’s solution (15: 15: 1; aqueous solution saturated with picric acid: 40% formalin: glacial acetic acid) at room temperature and flow rate 0.5 ml/min for P0 and 00.7 ml/min for P21 using an infusion pump (Model 07525-40, Masterflex, Gelsenkirchen, Germany). The brain was removed and immersed for 3 days in Boüin’s solution and 30% sucrose solution for three additional days. Once fixed and cryoprotected, the brain was frozen, and coronal slices (10 μm thick) were obtained with a cryostat (Leica CM1850 UV, Wetzlar, Germany) and recovered on slides treated with poly-L-lysine (Sigma-Aldrich).

Four coronal slices (10 μm thick) containing the M1 were recovered every 30 μm containing slices at ∼1–0.4 mm from Bregma at P0 (Figure 1A), and ∼1.0–0.6 mm for P21, according to Khazipov et al. (2015) (Figure 2A).

The sections were rinsed with PBS and incubated for 1 h with the blocking and permeabilization solution (PBS with 10% normal goat serum and 0.3% Triton X-100). The autofluorescence was blocked with TrueBlack (Biotium, Hayward, CA, United States), and the tissue was incubated overnight at 4°C with primary antibodies (Table 1) diluted in PBS containing 10% normal goat serum. The following day the tissue was washed three times (10 min each) with PBS at room temperature and then incubated for 1 h at room temperature with the corresponding secondary antibodies (Table 1). Nuclei were stained with DAPI (4′, 6-diamidino-2-phenylindole; 1 μg/ml) for 10 min. After washing three times for 10 min with PBS, the tissue was mounted with Aqua Poly/mount (Polyscience, Warrington, PA, United States).

Micrographs of M1 were obtained with a charge-coupled device camera (ORCA-Flash 2.8 CCD model C11440; Hamamatsu, Honshu, Japan) coupled to the epifluorescence microscope (Olympus IX-81, Shinjuku, Tokyo, Japan). Five (P0) or seven (P21) focal planes from the pial surface to the lateral ventricle were captured per slice (10×) to reconstruct the entire M1 area using Adobe Photoshop software (C6S; San Jose, CA, United States). The exposition and gain values used for the first micrograph from a control sample for each marker were maintained for all subsequent acquisitions. Fluorescence quantification was performed with Fiji software (Schindelin et al., 2012), and data were expressed as percentage of the fluorescence obtained in control samples.

The frontal cortices of both hemispheres were dissected from one P0 pup from different litters to obtain total protein extracts (Figure 1A). Three to five different animals from different litters per group (n = 3–5 different litters) were used.

The tissue was homogenized in lysis solution (25 mM Tris-HCl, pH 7.4, 1% IGEPAL, 100 mM NaCl) containing protease inhibitors (Amresco, Solon, OH, United States), the lysate was centrifuged at 13,000 × g for 10 min at 4°C, the supernatant was recovered, and the protein content was quantified by the Bradford method (Sigma-Aldrich; Bradford, 1976).

For electrophoresis, 20–80 μg of protein per sample and 2.5 μl of the molecular weight marker (Precision Plus Protein™ WesternC™ Blotting Standards, Bio-Rad or Trident Prestained Protein Ladder, GeneTex) were loaded on 10% polyacrylamide (SDS-PAGE) gels and proteins separated for 30 min at 70 V (4% separator gel) and 2 h at 100 V (10% concentrator gel), using the MiniProtean II system (Bio-Rad, Hercules, CA, United States). Proteins were then transferred to nitrocellulose membranes (Amersham™ Hybond™-ELC, Buckinghamshire, United Kingdom), using the Trans-Blot^® semi-wet transfer system (Bio-Rad; Villanueva, 2008) at 1.3 A/25 V for 12 min.

Non-specific binding of antibodies was blocked by incubating (1 h) with 7% fatty acid-free milk (SVELTY^® Milk, Nestlé, Mexico City, Mexico) in TBS buffer (20 mM Tris-base, 150 mM NaCl, pH 7.4). Membranes were incubated overnight at 4°C with the primary antibodies (Table 1) diluted in TBS, washed with TBS-T solution (TBS containing 0.1% Tween-20), and then incubated with secondary antibodies (Table 1) for 1 h at room temperature. Bands corresponding to the proteins of interest were detected by scanning the membranes on the Odyssey Clx system and analyzed with the Image Studio software version 4.0 (Li-COR Bioscience, Lincoln, NE, United States). The fluorescence signals obtained for GAPDH or α-actin were used as internal controls.

Golgi-Cox staining was performed to evaluate the structure and orientation of pyramidal neurons in the M1 from P21 animals (FD Rapid GolgiStain Kit, FD Neurotechnologies, Columbia, MD, United States). Brains from 3 or 4 animals from different litters (n = 3–4) were obtained after decapitation, washed with PBS at 4°C, and after removal of blood vessels and meninges cut into two coronal segments placed in the impregnation solution for 12 h, and stored protected from the light in a new impregnation solution for two weeks at room temperature. Tissues were then transferred to a cryoprotective solution for three days at 4°C.

Coronal sections were obtained with a cryostat at 60 μm thickness to improve staining, development, and washing efficacy. Sections were placed in the developing solution for 1 min, washed in double-distilled water for 5 min, dehydrated in alcohol (50, 75, 96, and 100%; 5 min in each dilution), transferred to xylol, and mounted with Entellan (Merck, Darmstadt, Germany).

For morphological analysis, micrographs (20×) at ∼1.0–0.6 mm from Bregma (Khazipov et al., 2015) of the pyramidal neurons from each M1 layer of both hemispheres were obtained from at least three subjects from three different litters per group. To reconstruct the dendritic arbor, only completely impregnated pyramidal neurons, whose dendritic spines and the dendritic arbor could be visualized in different planes, were documented. At least eight neurons per layer (I–II, III–IV, and V–Vl) from three different subjects of different litters per group were analyzed.

For dendritic arbor analysis, a printed image was obtained from the reconstruction of a series of micrographs. Then, dendritic arborization was reconstructed using a magnifying glass (0.25×), a fine-point black marker, and a white sheet placed on each impression by drawing the dendrites from different planes into a single plane. Finally, drawings were scanned to perform the Sholl analysis (Sholl, 1953) and dendritic complexity analysis, using the Fiji software (Schindelin et al., 2012).

For the Sholl analysis, concentric circles were drawn every 10 μM from the soma, and the number of intersections between the increasing circles and the dendritic arbor was quantified. The dendritic complexity index (DCI) was determined by assigning an ordinal value to each of the dendrites, referring to the order in which a branch appears in relation to the first branch (primary dendrite), and applying the following formula (Chameau et al., 2009):

Two P21 rats from three different litters (n = 3) for a total of six animals were used for electrophysiological experiments. Animals were deeply anesthetized with sodium pentobarbital (50 mg/kg, ip) and rapidly decapitated. The brain was quickly removed and placed in an ice-cold sucrose solution saturated with carbogen (95% O₂/5% CO₂) containing (in mM): sucrose 210, KCl 2.8, MgSO₄ 2, Na₂HPO₄ 1.25, NaHCO₃ 26, D-(+)-glucose 10, MgCl₂ 1, and CaCl₂ 1, pH 7.2–7.35. After 30–45 s, the hemispheres were separated along the mid-sagittal line, and coronal slices (385 μm thick) containing the M1 were obtained with a vibratome (VT1000S; Leica, Nussloch, Germany). Slices were incubated at 34°C for 30 min and at least for 60 min at room temperature in an incubation solution containing (in mM): NaCl 125, KCl 2.5, Na₂HPO₄ 1.25, NaHCO₃ 26, MgCl₂ 4, D-(+)-glucose 10, and CaCl₂ 1, pH 7.4.

Individual slices were transferred to a submersion recording chamber and placed on an upright Nikon FN1 microscope (Nikon Corporation, Minato, Tokyo, Japan). The slices were continuously perfused with artificial cerebrospinal fluid (aCSF) at a constant flow rate (2-3 ml/min) with a peristaltic pump (120S, Watson-Marlow, Wilmington, MA, United States). The aCSF composition was (in mM): NaCl 125, KCl 2.5, Na₂HPO₄ 1.25, NaHCO₃ 26, MgCl₂ 2, D-(+)-glucose 10, and CaCl₂ 2; pH 7.4, saturated with carbogen.

Coronal slices containing pyramidal neurons from layers V/VI of M1 were visualized using a Nikon FN-S2N microscope (Nikon, Tokyo, Japan) coupled to an infrared differential interference contrast coupled camera. Borosilicate pipettes were obtained with a horizontal puller (Flaming-Brown P-97, Novato, CA, United States) with a final resistance of 3–6 MΩ when filled with an intracellular solution containing (in mM): potassium gluconate 135, KCl 10, EGTA 1, HEPES 10, Mg-ATP 2, NA₂-GTP 0.4, and phosphocreatine 10; pH 7.2–7.3 and 290–300 mOsm/L). Whole-cell, patch-clamp recordings were performed with an Axopatch 200B amplifier (Molecular Devices, San José, CA, United States), digitized at a sampling rate of 10 kHz, and filtered at 5 kHz with a Digidata 1322A (Axon Instruments, Palo Alto, CA, United States). Electrophysiological signals were acquired and analyzed offline with pCLAMP10.6 software (Molecular Devices).

To analyze intrinsic membrane properties and neuronal excitability, current-clamp experiments were performed under basal conditions and in the presence of HA (1, 10, and 100 μM). Resting membrane potential (RMP) was determined immediately after the initial break-in from gigaseal to whole-cell configuration. Once RMP was stabilized, a series of negative and positive current steps from −300 pA to rheobase (1 s duration, 30 pA increments) were injected to determine input resistance (Rin), membrane time constant (τ), and the rheobase current. The input resistance (Rin) was determined as the slope of a linear function fitted to the current–voltage (I–V) relationship near the RMP. The membrane time constant (τ) was determined by fitting an exponential function to a voltage response elicited by the injection of a pulse of negative current (−30 pA). The amplitude of the “sag” conductance, defined as the voltage drop in response to a hyperpolarizing current step, was determined by injecting a pulse of −300 pA. Firing frequency was evaluated by applying current pulses from 0 to 500 pA (1 s duration, 50 pA increments) and quantifying the number of evoked action potentials. The rheobase was calculated as the current required to evoke at least one action potential.

Four P21 rats from three different litters (n = 3) per group were used to evaluate the effect of the H₁R antagonist/inverse agonist mepyramine (1 μM) or the H₂R antagonist/inverse agonist tiotidine (1 μM) on firing frequency in response to the injection of a current step of 300 pA (1 s). Each neuron was registered under the following conditions: (a) no drugs (basal); (b) in the presence of mepyramine or tiotidine; and (c) in the presence of mepyramine or tiotidine and 10 μM HA.

For the electrophysiology experiments, a maximum of two slices per animal and only one neuron per slice were recorded to avoid biased interpretations.

Glutamate release was performed as previously reported (Giuliani et al., 2011; Casas et al., 2013) in cross-chopped P21 striatal slices (250 × 250 μm) obtained from four pups from the same litter for a total of seven litters per group (n = 7). Striatal slices were placed in ice-cold Krebs-Henseleit (KH) solution containing (in mM): NaCI 126, KCI 3, MgSO₄ 1, KH₂PO₄ 1.2, NaHCO₃ 25, D-glucose 11, pH 7.4 after saturation with O₂/CO₂, 95:5% v:v). CaCl₂ was omitted to reduce excitotoxicity.

Thereafter, slices were equilibrated in KH solution containing 1.8 mM CaCl₂ for 30 min at 37°C, changing the solution every 10 min. Slices were then transferred to 1 ml KH solution containing 100 nM [³H]-glutamate (Perkin Elmer, Boston, MA, United States) in the presence of dihydrokainic acid and aminooxyacetic acid (200 μM each) to prevent [³H]-glutamate uptake by glial cells and degradation by transaminases, respectively.

After loading, slices were washed three times with KH solution and randomly distributed (30 μl of sedimented slices) in a perfusion chamber system (Brandel 2500, Gaithersburg, MD, United States) and perfused (1 ml/min) with KH solution for 20 min. Two basal fractions were collected (1 ml each), and [³H]-glutamate release was stimulated at the third fraction by switching to a KH solution containing 50 mM KCl (equimolarly substituted for NaCl). The perfusion was returned to standard KH solution and seven fractions were further collected. The Ca²⁺-dependence of [³H]-glutamate release was tested in KH solution without CaCI₂ added.

At the end of the perfusion, the tissue was recovered and treated with 1 ml HCI (1 N) for 1 h, and the tritium content was determined by scintillation counting. Fractional release values were normalized to the fraction collected immediately before fraction 3, and the area under the curve was obtained to determine total [³H]-glutamate release.

Histamine content was determined by ELISA (cat. 17-HISRT-E01-RES, sensitivity 0.2 ng/ml, 100% HA specificity). The frontal cerebral cortex (∼2.8 to −0.4 mm from Bregma; Khazipov et al., 2015) from one animal of different litters for a total of 5–9 litters per group (n = 5-9).

Following the protocol recommended by the supplier (ALPCO R immunoassays, Salem, NH, United States), the tissue was homogenized in 100 μl ice-cold PBS (Polytron PT 2100 Homogenizer, Kinematica, Luzern, Switzerland), centrifuged (10,500 × g for 5 min), and the supernatant was recovered to determine per duplicate the HA content using a reference curve constructed with 0, 0.5, 1.5, 5, 15, and 50 ng/ml HA and by determining the absorbance at 450 nm in a multiple detection system (Glomax, Promega, Madison, WI, United States). Values were corrected by protein content determined by the Bradford method (Bradford, 1976).

H₁R blockade by the systemic or intrauterine administration of chlorpheniramine at E12 leads to a reduction in FOXP2⁺ deep-layer neurons in the dorsal telencephalon at E14. The cells that preferentially express the H₁R in the developing cortex are NSCs located in the ventricular zone (Supplementary Figure 1). As a first approach, we determined the expression of Foxp2 together with two other cortical markers (Satb2 and Tbr1) by qRT-PCR. We found a reduction in the relative expression of Satb2 and Foxp2 to 21 and 56% of control values, respectively, in the frontal cerebral cortex of neonates from chlorpheniramine-treated rats, while Tbr1 relative expression was not affected (Figure 1B).

WB analysis confirmed the lack of effect on TBR1, as the protein level analysis did not show any significant difference with the control group. In contrast to qRT-PCR analysis, this approach showed increased SATB2 and FOXP2 levels in the frontal cortex of P0 neonates of chlorpheniramine-treated rats (240 and 250% of control neonates, respectively; Figure 1C). In accordance with WB, the global immunofluorescence analysis (from the pial to the deepest layer) in M1 showed that there was no effect on TBR1, whereas the fluorescence signals for SATB2 and FOXP2 increased to 180 and 340% of control values, respectively (Figure 1D).

For the layer distribution analysis, the cortex was divided into upper (I and II-IV) and deeper (V/VI) layers, taking into account that layers V/VI represent ∼50% of the total area in the motor cortex of the rat. Our results showed that in neonates of the control group, SATB2 was only expressed in M1 layer I, while FOXP2 and TBR1 were expressed in both upper and deeper layers (Figure 1D), as previously reported for P0 rats (Valle-Bautista et al., 2020). Indeed, the transcription factor immunostaining after birth in the rat is different from the adult cortex and other cortical areas in P0 mice (Huang et al., 2013; Srivatsa et al., 2014). Furthermore, SATB2 did not show the ‘classical’ nuclear immunostaining described for transcription factors (Figure 1D).

The offspring exposed to chlorpheniramine during embryo development showed changes in the distribution of SATB2 and FoxP2, whose location extended ventrally to layer III and throughout all layers, respectively (Figure 1D).

In P21 pups, the layers were identified by their marker distribution in the control animals, as previously reported (Watakabe et al., 2012; Kast et al., 2019). We took as reference the FOXP2 mark to delimit layers V/VI, and SATB2 for upper layers. Although layer IV in the M1 mouse is not visually appreciated, it is present in the rat, but is thinner than in other cortical areas. Hence, we consider this layer a SATB2 expressing area as reported in the literature (Skoglund et al., 1997; Kast et al., 2019; Valle-Bautista et al., 2020). The SATB2, FOXP2, and TBR1 immunostaining were localized as expected in the control group (Figure 2B). In contrast, in pups from chlorpheniramine-treated rats, SATB2 neurons were preferentially observed in layers V and VI. In turn, FOXP2 and TBR1 transcription factors were dorsalized, with FOXP2 located from layers II to VI and TBR1 from layers IV to VI (Figure 2B). Furthermore, in the offspring of chlorpheniramine-treated rats, the total fluorescence increased significantly to 177, 178, and 141% of the control group for SATB2, FOXP2, and TBR1, respectively (Figure 2C).

To analyze the morphology of M1 pyramidal neurons in P21 pups, reconstructions of the neurons stained with the Golgi-Cox were performed (Figure 3A). Compared to the control group, the systemic administration of chlorpheniramine from E12 to E14 promoted significant changes exclusively in deep-layer neurons of P21 pups, with a significant reduction in the number of proximal processes at ∼60 and ∼80 μM from the soma, as well as in the DCI (Figures 3B,C).

Since morphological changes were observed in M1 deeper layers, we determined the electrophysiological properties in pyramidal neurons from these layers. Under basal conditions, no differences between groups were observed in the neuronal passive and active electrophysiological properties (Figures 4–6). However, while HA increased the membrane resistance (Rin) in the control group, in the chlorpheniramine group this parameter was significantly lower compared to the control group and was not affected by HA (Table 2).

In basal conditions, the firing frequency did not differ between groups (Figures 5A,B). However, and as expected, HA (1, 10, and 100 μM) increased the firing frequency in neurons from the control group, with significant effect from 150 pA and at 10 and 100 μM (Figure 5C). In contrast, HA did not alter the firing frequency of neurons from the chlorpheniramine group (Figure 5D).

Phase plots from the first and tenth action potentials elicited by a current injection of 300 pA revealed that under basal conditions the conductances underlying the depolarization (Na⁺) and repolarization (K⁺) phases of the action potential were not different in neurons from the offspring of chlorpheniramine-treated rats. However, in the presence of 10 μM HA, the maximum rate of the depolarizing phase increased in these neurons (Figure 6), suggesting changes in either the expression or the biophysical properties of fast-inactivating Na⁺ currents.

HA increases neuronal excitability through H₁R activation (McCormick and Williamson, 1991; Ji et al., 2018; Zhuang et al., 2018). Therefore, H₁R density was determined in P21 frontal cortex membranes from both groups. Unexpectedly, the products of chlorpheniramine-treated dams showed increased H₁R density compared to the control group (69 ± 7 and 42 ± 7 fmol/mg protein, for the control and chlorpheniramine groups, respectively; P = 0.0046; t = 3.74, df = 9, unpaired t-test; control n = 6 and chlorpheniramine n = 5). Likewise, an increase in cortical HA content was observed (826 ± 278 and 260 ± 30 ng/mg protein for the control and chlorpheniramine groups, respectively; P = 0.0170; t = 2.76, df = 12, unpaired t-test; control n = 9 and chlorpheniramine n = 5).

The increase in H₁R density and HA content in pups from chlorpheniramine-treated rats is counterintuitive with the decrease in HA-induced excitability. Moreover, the increase in neuronal firing observed in neurons from animals of the control group could also involve H₂R activation (Panula et al., 2015). Therefore, the participation of each receptor was evaluated by determining the firing frequency at a depolarizing pulse of 300 pA in the presence of the H₁R antagonist/inverse agonist mepyramine (1 μM) or the H₂R antagonist/inverse agonist tiotidine (1 μM) and HA (10 μM).

In neurons from animals of the control group, mepyramine decreased basal firing frequency from 24.80 ± 3.72 to 20.20 ± 3.05 Hz and prevented the increase in firing induced by HA (20.00 ± 2.66 Hz). In contrast, tiotidine had no effect on its own but prevented the HA-induced increase in firing frequency (18.75 ± 2.23, 19.5 ± 2.07, and 21 ± 1.87 Hz for basal, tiotidine, and tiotidine + HA, respectively; Figures 7A,B). In neurons from animals of the chlorpheniramine group, mepyramine did not modify the basal firing frequency (21.67 ± 3.24, 21.83 ± 2.76, and 20.67 ± 2.81 Hz in basal, mepyramine, and mepyramine + HA conditions, respectively). However, it prevented the HA effect for basal, tiotidine, and tiotidine + HA conditions (12.17 ± 3.26, 15.17 ± 3.42, and 14.67 ± 3.98, respectively; Figures 7A,B). Tiotidine alone promoted a significant increase in the firing frequency in the basal condition and prevented the HA-induced increase (Figures 7A,B).

To further evaluate the functionality of M1 deep cortical neurons, which project to the striatum, K⁺-evoked [³H]-glutamate release from striatal slices was evaluated. The release of the labeled neurotransmitter evoked by 50 mM K⁺ was markedly dependent on the presence of Ca²⁺ ions in the perfusion medium (84.3 ± 5.6% of total release, P < 0.0001; t = 18.80, df = 8, unpaired Student’s t-test; five experiments; Figures 8A,B). K⁺-evoked [³H]-glutamate release was significantly lower in slices from the offspring of chlorpheniramine-treated rats (72.7 ± 7.3% of the release in slices from animals of the control group; Figures 8C,D).