• hypoglossal nucleus;
  • N-methyl-d-aspartate receptor kinetics;
  • nucleus tractus solitarius;
  • pre-Bötzinger complex;
  • rat;
  • reverse transcription–polymerase chain reaction


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

NMDA receptors are involved in a variety of brainstem functions. The excitatory postsynaptic NMDA currents of pre-Bötzinger complex interneurons and hypoglossal motoneurons, which are located in the medulla oblongata, show remarkably fast deactivation kinetics of approximately 30 ms compared with NMDA receptors in other types of neurons. Because structural heterogeneity might be the basis for physiological properties, we examined the expression of six NMDA receptor subunits (NMDAR1, NR2A−2D, and NR3A) plus eight NMDR1 splice variants in pre-Bötzinger complex, hypoglossal and, for comparison, neurons from the nucleus of the solitary tract in young rats using single cell multiplex RT–PCR. Expression of NR2A, NR2B, and NR2D was observed in all three cell types while NR3A was much more abundant in pre-Bötzinger complex interneurons, which belong to the rhythm generator of respiratory activity. In hypoglossal neurons, the NMDAR1 splice variants NMDAR1–4a and NMDAR1–4b were found. In neurons of the nucleus of the solitary tract, instead of NMDAR1–4b, the NMDAR1–2a splice variant was detected. This differential expression of modulatory splice variants might be the molecular basis for the characteristic functional properties of NMDA receptors, as neurons expressing a special NMDAR1 splice variant at the mRNA level show fast kinetics compared with neurons lacking this splice variant.

Abbreviations used

a-amino-5-phosphonovaleric acid


excitatory postsynaptic current


nucleus hypoglossus


NMDA receptor


nucleus tractus solitarius


pre-Bötzinger complex



Ionotropic glutamate receptors are found throughout the mammalian brain, where they constitute the major excitatory neurotransmitter system. In spontaneous oscillating networks like the mammalian respiratory network, excitatory glutamatergic signals are of essential importance during generation and maintenance of rhythmic activity (Pierrefiche et al. 1994; Bonham 1995). A variety of ionotropic glutamate receptors, including NMDA receptors, are expressed in respiratory-related neurons. The NMDA receptor subfamily comprises three different types of subunits, namely NR1, NR2 and NR3 subunits. NMDAR1 subunits show RNA splicing at three independent positions, giving rise to eight different splice variants, NMDAR1–1a to NMDAR1–4a, and NMDAR1–1b to NMDAR1–4b (Hollmann et al. 1993). These splice variants differ in their potentiation by protein kinase C (Durand et al. 1993), zinc (Hollmann et al. 1993), spermine (Zheng et al. 1994) and neurosteroids (Malayev et al. 1998; Ceccon et al. 2001). Co-expression of NMDAR1 with any of the four NR2 subunits (NR2A to NR2D) yields much larger currents than NMDAR1 alone (Ikeda et al. 1992; Kutsuwada et al. 1992). Assuming a tetrameric subunit stoichiometry, a functional NMDA receptor can be built by combining two glycine-binding NMDAR1 subunits and two glutamate-binding NR2 subunits (Laube et al. 1998). NR3A and NR3B, recently cloned modulatory subunits, have the opposite effect compared with NR2 and decrease currents when incorporated in heteromeric receptors (Sucher et al. 1995; Nishi et al. 2001; Matsuda et al. 2002). Receptors containing both NR1 and NR3, but not NR2, subunits in vitro can form excitatory glycine receptors (Chatterton et al. 2002), although it is not clear if this occurs in neurons (Matsuda et al. 2003). The ratio of incorporation of NR3A in a functional NMDA receptor complex remains to be determined. For a review of NMDA receptors, see Cull-Candy et al. (2001).

NMDA receptor-evoked currents play a critical role in essential brainstem functions such as the modulation of the respiratory rhythm and the movement of the tongue. Rhythmic tongue movement during suckling behaviour is probably represented by NMDA-elicited rhythmic activity in the hypoglossal nucleus (NH) (Katakura et al. 1995). Cerebral blood flow is modulated by NMDA receptors of the nucleus of the solitary tract (NTS) (Morino et al. 1994). NMDA receptors expressed in neurons of the pre-Bötzinger complex (PBC) have a regulatory function in respiratory rhythm generation (Otsuka et al. 1994). This modulatory role for the NMDA receptors is also indicated by the finding that mice lacking the NMDAR1 gene die shortly after birth as a result of respiratory failure (Forrest et al. 1994). Spontaneous NMDA currents are a property of the network and entirely independent of the spontaneous rhythmic-respiratory activity which is triggered by rhythmic oscillations of the endogenous pacemaker neurons. Those rhythmic oscillations are determined by the ratio of Na+ currents and leak currents in the pacemaker neurons (Del Negro et al. 2002).

A prerequisite for the determination of the functional role of NMDA receptors in respiratory rhythm generation is a detailed knowledge of the electrophysiological and molecular biological properties of NMDA receptors expressed in the cells involved. To date, kinetics and subunit composition, which determine the properties of the NMDA receptors, have not been investigated in the respiratory pacemaker system, the PBC. In the present study we analyzed the maximal breathing frequency of young rats during hyperpnoea, as well as the kinetics of the NMDA EPSCs and the NMDA receptor subunit composition of single PBC interneurons and NH motorneuons in comparison with NTS interneurons which have been investigated previously (Titz and Keller 1997).

Materials and methods

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Determination of maximal breathing frequency in juvenile rats

Breathing rates of young rats were determined by measuring pressure transients as previously described (Jacquin et al. 1996). Juvenile rats were placed in a chamber (20 mL) connected to a differential pressure transducer (model DP103-14, Validyne Engineering, Northridge, CA, USA). Normal breathing rates were obtained using constant temperature at 31°C while pressure was measured with reference to a second chamber of identical volume. Breathing rates were recorded using EPC-9 software and an ITC−16 interface (Heka Elektronic, Lambrecht, Germany) or Maclab hard- and software (AD Instruments, Colorado Springs, CO, USA). To determine the maximal breathing frequency, rats were stressed by placement in an unfamiliar environment where they immediately start searching for a comfortable place. After a few minutes observing this behaviour, animals were placed back into the recording chamber and breathing frequency measurements were started instantly. Breathing rates were elevated in comparison with those of unstressed control animals.

Preparation of brainstem slices

Brainstem slices were cut from isolated brains of 6–11 day-old Wistar rats. Under deep ether anaesthesia, rats were decapitated and the brains were isolated from the skull. The isolated brainstem was put in cold artificial cerebrospinal fluid (aCSF: 118 mm NaCl, 3 mm KCl, 1 mm MgCl2, 25 mm NaHCO3, 1 mm NaH2PO4, 1.5 mm CaCl2, 30 mm glucose) bubbled with carbogen (95% O2/5% CO2; pH at 7.4). The brainstem was glued to an agar block with the rostral side up, mounted in an upright position with its rostral side upwards and the dorsal side facing the slicer blade. The brainstem and the agar block were inclined 20° in the dorsal direction and transverse slices of 150–200-μm thickness were cut. Characteristic topographic ‘landmarks’ including the facial nucleus, inferior olive, nucleus of the solitary tract (NTS), hypoglossal nucleus (NH) and the area of the pre-Bötzinger complex (PBC) were used to yield a reproducible cutting plane (Fig. 1). After a 20-min period of recovery in aCSF at room temperature (21°C), the slices were transferred to the recording chamber. The flow rate was adjusted to 5–6 mL/min for an adequate oxygen supply. By using infrared differential interference contrast optics (Dodt and Zieglgansberger 1994) individual cells could be visualized by infrared-sensitive video equipment. Neurons were then immediately harvested for single-cell PCR experiments (Lambolez et al. 1992). Harvesting included the establishment of the whole cell patch-clamp configuration and suction of the cytoplasm into the pipette. Pipettes were pulled from cleaned borosilicate glass capillaries (Kimax-51, Kimble, Vineland, NJ, USA) using a vertical puller (DMZ-Puller, Zeitz-Instrumente, Augsburg, Germany) which had outer diameters of 2–3 μm and DC resistances of 1.5–2.5 MΩ using a special intracellular standard solution described elsewhere (Frermann et al. 1999). The solution in the pipette contained a mixture of 2 μL 5 × reverse transcription buffer (Gibco BRL, Karlsruhe, Germany 250 mm Tris pH 8.3, 375 mm KCl, 15 mm MgCl2) and 4.5 μL H2O. All experiments were carried out in accordance with the guidelines of the Ethics Committee of the Medical Faculty of the University of Göttingen.


Figure 1. Identification of different brainstem nuclei in the acute slice preparation. A schematic drawing of transversal sections showing different respiratory-related regions of the medulla oblongata in rats is depicted. Experiments were carried out in optically identified kernels of the brainstem.

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Measurements of NMDA EPSCs

Patch-clamp experiments were performed using slice preparations as previously described (Titz and Keller 1997). Pipettes were filled with an ‘intracellular’ solution containing (in mM) 140 KCl or 140 CsCl mm, 2 MgCl2, 10 Hepes, 2 ATP, 0.4 GTP, 10 EGTA, 1 CaCl2; Osm 316, pH: 7.2). Patch pipettes were pulled from ethanol-cleaned autoclaved borosilicate glass tubing (Hilgenberg, Malsfeld, Germany) and were heat polished before use. The tip diameters were 1–3 μm. Neurons were held in the whole cell voltage-clamp configuration at −70 to −60 mV using a standard patch-clamp amplifier (EPC-9, Heka, Lambrecht, Germany). Series resistances were higher than 15 MΩ. No compensation for liquid junction potentials was performed. Unless stated otherwise, whole cell measurements were recorded with sampling frequencies of 100 kHz and filtered (4-pole Bessel filter 2.9 kHz) before analysis. NMDA receptor-evoked EPSCs in single patch-clamped neurons were pharmacologically isolated using extracellular application of bicuculline, strychnine and CNQX (10 μm each; purchased from Sigma, Munich Germany). To raise the spontaneous glutamate release extracellular potassium concentrations were transiently elevated (4–6 mm). In hypoglossal neurons, application of serotonin (5–10 μm) was also used to increase the spontaneous activity. The cells analyzed by RT-PCR were not the same as those recorded electrophysiologically. Therefore, one has to be careful in correlating both kinds of data. The cells analyzed in the three nuclei were homogeneous in their electrophysiological properties, and cells for both types of experiments were picked by the same criteria.

Harvesting of single neurons and reverse transcription

Single cells of the optically identified kernels of the PBC, the NH and the NTS were whole-cell patch-clamped and immediately aspirated into the pipette. Tips of the patch-clamp pipettes were broken at the bottom of a siliconized tube and the contents expelled. A solution was added containing hexamer random primers (Roche, Mannheim, Germany; final concentration 12.5 μm), four deoxynucleotide triphosphates (Pharmacia, Freiburg, Germany; 0.5 mm final concentration each), dithiothreitol (BRL, 10 mm final concentration), 20 U of ribonuclease inhibitor (Promega, Madison, WI, USA) and 100 U of Moloney murine leukaemia virus reverse transcriptase (BRL). The resulting 10-μL mix was incubated for 1 h at 37°C for synthesis of single-stranded cDNA. The entire reaction was used as template for a multiplex PCR.

Multiplex PCR

Following reverse transcription, the NMDA receptor subunits were amplified. β-Actin was used as an internal housekeeping gene control (see Fig. 7a). The primers are listed in Table 1.


Figure 7. Control experiments for single cell PCR. The housekeeping gene actin was used as an internal control for the presence of RNA in the samples (a). Only neurons in which actin was detectable were used in this study. PCR products were analyzed on ethidium bromide-stained agarose gels. Reagent controls represent contamination checks for reagents used in harvesting the neurons and in the subsequent RT–PCR, and were performed in parallel to the analyzed neurons. Rat brain cDNA was used as a control for failure of the PCR reaction. To confirm the results of the PCR, the PCR products were subjected to restriction analysis. Examples are shown for NMDAR1-a/NMDAR1- b (b), NMDAR1-1/NMDAR1-2 (c), NMDAR1-3/4 (d), NR2B (e), NR2D (f) and NR3A (g).

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Table 1.  PCR primers
Gene/splice variantSequence (from 5′-to 3′)

To analyze NMDAR1 splice variants, three primer pairs were designed, the first to amplify both N-terminal (NMDAR1-a and NMDAR1-b) variants, a second for one pair of C-terminal variants (NMDAR1-1 and NMDAR1-2) and a third primer pair for the other two C-terminal variants (NMDAR1-3 and NMDAR1-4). Although the primer binding sites used for NMDAR1-3 and NMDAR1-4 are present in all NMDAR1 splice variants, under the experimental conditions used PCR products were obtained only for NMDAR1-3 and NMDAR1-4, but not for the other C-terminal splice variants.

Because of the use of single cell PCR, real-time techniques for mRNA quantification were not applicable. To the RT reaction, 5 nmol of each deoxyribonucleotide triphosphate (Pharmacia), 2.5 U of Taq DNA polymerase (Hilden, Germany) and buffer supplied by the manufacturer were added to a final volume of 80 μL. Two drops of mineral oil were used as overlay and after 30 s at 85°C in the pre-heated thermocycler (MJ Research PTC 100), 20 μL containing the primer pairs (10 pmol each) were added. The initial denaturation (3 min at 94°C) was followed by 20 cycles (94°C/30 s, 60°C/30 s, 72°C/35 s) of PCR and a final elongation of 5 min at 72°C. Two microlitres of the first PCR product were used as a template in a second round of PCR amplification. In this second round, each subunit was amplified individually using its specific primer pair and the same PCR program as for multiplex PCR, but with 35 cycles instead of 20. Twenty cycles were employed in the first round to obtain enough material to enable analysis of all different splice variants and genes, but at the same time to minimize interference between the many different primers and PCR products. Thirty-five cycles in the second PCR round were applied to reach maximal sensitivity. Ten microlitres of each amplification product were run on a 2% agarose gel in parallel with a molecular weight marker (100 bp ladder, MBI, St. Leon-Rot, Germany), and stained with ethidium bromide. As a positive control for the multiplex PCR, 25 ng of reverse-transcribed rat brain RNA was used as template. The predicted sizes of the PCR-generated fragments were (in bp): 398 (NMDAR1-a), 461 (NMDAR1-b), 488 (NMDAR1-1), 377 (NMDAR1-2), 433 (NMDAR1-3), 322 (NMDAR1-4), 257 (NR2A), 314 (NR2B), 464 (NR2C), 265 (NR2D), 417 (NR3A), and 255 (actin).

Restriction analysis

To confirm the results of the agarose gel electrophoresis, the PCR products were cut with sequence-specific restriction enzymes. Briefly, the remaining 90 μL of the second PCR products were extracted once with chloroform/isoamylalcohol (24 : 1), ethanol-precipitated and finally dissolved in 20 μL H2O. The purified PCR products were cut with appropriate restriction enzymes and analyzed by agarose gel electrophoresis as described above. The predicted sizes and the corresponding restriction enzymes were (in bp): 95/303 (PstI, NMDAR1-a), 95/366 (PstI, NMDAR1-b), 217/271 (PstI, NMDAR1-1), 106/271 (PstI, NMDAR1-2), 162/271 (PstI, NMDAR1-3), 51/271 (PstI, NMDAR1-4), 44/213 (MscI, NR2A), 55/259 (BclI, NR2B), 229/235 (Eco47III, NR2C), 107/158 (DrdI, NR2D), and 144/273 (XcmI, NR3A). Examples are shown in Fig. 7(b–f).

Statistical analysis

Statistical significance of differences between cell types for a given subunit was calculated using Fisher's exact test if the number of analyzed cells was below 15, and otherwise using the ‘four arrays test’ (Sachs 1974).


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Measurements of breathing rate

The breathing frequency in young rats is known to be high. To determine the exact frequency for normal breathing and hyperpnoea we measured the in vivo breathing rhythm from animals between 4 and 12 days prior to electrophysiological measurements. A plethysmographic device to monitor animal ventilation was used (Jacquin et al. 1996). Resting breathing frequencies and breathing under physical stress were measured. Normal breathing frequencies were very constant in individual animals. The absolute frequency had values around 3.5 Hz with a standard deviation of less than 10% in individual animals. Breathing rhythms between individuals of similar age showed larger variations. At the age of 5 days, animals showed breathing frequencies between 2.8 and 4.2 Hz (average 3.5 Hz ± 0.64, n = 8) depending on the fitness and the metabolic condition of each individual. Average periods between two maximal inspiratory peaks had absolute values of 307 ms (± 35.8, n = 11). During physical stress, breathing rates were elevated up to inspiratory periods below 200 ms (≈ 5 Hz). Older animals showed slightly lower breathing frequencies (with a period of c. 350 ms). In Fig. 2, the breathing frequency of a 5-day-old animal is depicted. In Fig. 2(c), the normal breathing frequency is shown whereas Fig. 2(a and b) shows the hyperpnoea under physical stress.


Figure 2. Normal and elevated in vivo breathing rhythms in juvenile rats. Examples of breathing measurements from 5-day-old rats are shown. (c) Shows the resting breathing frequency, (b) elevated frequency and (a) breathing frequency under massive physical stress (hyperpnoea) measured in juvenile rats. (a, b) Irregularities concerning the baseline are the result of head movements because of the stress situation.

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Spontaneous respiratory-related rhythmic activity in single cells

Electric activity was measured during spontaneous respiratory bursts in single interneurons of the pre-Bötzinger complex. Electric activity of single neurons is highly correlated with extracellularly recorded activity of effector nerves like the nervus phrenicus or the nervus hypoglossus (Smith et al. 1990; Frermann et al. 1999).

Single cell activity of the pre-Bötzinger complex was measured using the current-clamp and voltage-clamp mode of the patch-clamp technique. In Fig. 3 spontaneous activity measured in two cells of the pre-Bötzinger complex is depicted. The upper trace shows rhythmic activity of one cell measured in the current-clamp mode; the lower trace was taken from another cell measured in the voltage-clamp mode. Typical respiratory-related electric oscillations could be recorded for several hours. Rhythmic activity in the in situ preparation was slower than the breathing rhythm observed in vivo. This has also been observed in other studies (Frermann et al. 1999) and is caused at least partially by the decreased temperature (29°C vs. 37°C). In single cells, bursts of rhythmic activity were elicited as a result of synchronous receptor activation stimulated by a multitude of neurons in the network. In the voltage-clamp mode action potentials during bursts could be suppressed using holding potentials of approximately −70 mV to −80 mV (data not shown). Postsynaptic activity revealed by inhibitory post synaptic currents (IPSCs) (inhibitory post synaptic currents) and EPSCs could not be suppressed by hyperpolarizing voltages. Glutamate receptor currents, namely NMDA currents, were pharmacologically isolated in single cells of the network and the kinetics were determined by fitting with one exponential function.


Figure 3. Spontaneous respiratory-related rhythmic activity in single neurons. (a) The spontaneous activity during respiratory oscillations in an interneuron of the pre-Bötzinger complex known as the rhythm generator for breathing is depicted. This measurement was performed using the current-clamp mode of the patch-clamp technique. Repetitive bursts of action potentials are elicited by depolarisations according to network activity. (b) Oscillatory activity of a different pre-Bötzinger complex neuron measured in the voltage-clamp configuration. At holding potentials of −40 mV, rhythmic-respiratory activity is reflected by repetitive bursts of excitatory postsynaptic currents (EPSCs) recorded in voltage-clamp mode.

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NMDA EPSCs in neurons related to the respiratory rhythm generator

Pharmacologically isolated NMDA EPSCs were measured using the whole-cell patch-clamp configuration. Spontaneous synaptic currents were weak in all observed nuclei. They showed various kinetics and amplitudes. Spontaneous currents showed amplitudes up to fivefold higher than those identified as NMDA currents (Figs 4a1/2 and b1/2). Some examples are shown in Figs 4(a and b). Pharmacological blockers against inhibitory synaptic currents (bicuculline and strychnine, 10 μm each) and against AMPA receptor-evoked currents (CNQX, 10 μm) were used to isolate NMDA currents. Holding potentials were set to −60 mV or +50 mV to inactivate a potentially persisting block of magnesium despite the use of magnesium-free solution (Fig. 4a) as effects from the remaining magnesium concentrations in the tissue have been reported (Konnerth et al. 1990). NMDA currents in the nucleus hypoglossus were small and fast, with amplitudes of 35.3 ± 15 pA and decay times of 36 ± 19 ms, n = 13 (Fig. 4a4). In the pre-Boetzinger complex, spontaneous activity was slightly lower (Fig. 4b1/2), but was also showing varying kinetics and amplitudes among animals. The amplitudes of NMDA currents had values of about 20 pA and decay times of 22 ± 12.2 ms, n = 5 (Fig. 4b2). Spontaneous activity of NMDA currents was very low but could be increased using fast application of depolarizing agents such as KCl (4–6 mm) or serotonin (5–10 μm). Kinetics of currents essentially remained the same; only the frequency of NMDA events was elevated. Figure 4(a3) shows repetitive occurrence of NMDA currents after short application of serotonin in hypoglossal neurons. Events identified as NMDA currents could be blocked reversibly by brief application of APV or permanently using 30 μm APV (Fig. 4a5). Decay times of EPSCs were determined by fitting a single exponential function to the EPSC. Even with blockage of inhibitory and AMPA-evoked currents, NMDA currents showed a spectrum of varying amplitudes, whereas the kinetics were fast in all observed EPSCs. The results are summarized in Table 2.


Figure 4. Time course of NMDA EPSCs in respiratory-related neurons. The time courses of pharmacologically isolated NMDA EPSCs of hypoglossal motoneurons (a) and interneurons of the pre-Bötzinger complex (b) were determined. (a) EPSCs of single hypoglossal motoneurons measured in the voltage-clamp mode of the patch-clamp technique are depicted. The frequency of spontaneous events in Mg2+-free solution was relatively high (1), whereas NMDA-elicited EPSCs were rare events (2). In some experiments, we used a permanent application of KCl or serotonin to stimulate cells in the environment of the measured cell and performed recordings at a holding potential of +50 mV (3). The kinetic of the currents was determined using single exponential functions (4). As a control, APV was applied to identify the NMDA currents (5). (b) Spontaneous activity and pharmacologically isolated NMDA EPSCs of neurons located in the pre-Bötzinger complex are shown. NMDA currents were less frequent than those observed in the hypoglossal nucleus (2), and their amplitudes were smaller. The time courses of NMDA currents were comparable (3). APV was used to verify the NMDA currents (traces not shown).

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Table 2.  Deactivation times of NMDA currents in different tissues
Neurons of the NTS320 ms Titz and Keller 1997
Hypoglossal motoneurons42 ms O'Brien et al. 1997
36 ms This paper
Interneurons of the PBCc. 22 ms This paper
Hippocampus granular cells50 ms240 msKeller et al. 1991
Hippocampal pyramidal cells66.5 ms353.9 msPerouansky and Yaari 1993
Hippocampal pyramidal cells12d: 81.6 ms853.8 msKirson and Yaari 1996
19d: 124.8 ms789.2 ms 
16w: 87.1 ms472 ms 
Interneurons in the CA1 field34.4 ms212.5 msPerouansky and Yaari 1993
Cerebellar granular cells53.6 ms216 msD'Angelo et al. 1994

Determination of NMDA receptor subunit composition

To investigate which NMDA receptor subunits are expressed in single neurons of the PBC, the NH and the NTS, these cell types were analyzed from 6- to 11-day-old rats using single cell RT–PCR. For most NMDA receptor subunits, 23–32 cells were analyzed per cell type. Exceptions were NR3A in the PBC (13 cells), NR3A in the NTS (6 cells) and NMDAR1-a in the NTS (12 cells).

NMDAR1-a splice variants were detected in 60–75% of the PBC (Fig. 5a), the NTS (Fig. 5b) and the NH neurons. Differences in detection rates of these subunits between the cell types were not statistically significant (Table 4).


Figure 5. Detection of NMDAR1 splice variants in single brainstem neurons. NMDAR1- a (398 bp) was detected in many neurons of PBC (a, lanes 36–41) and NTS (b, lanes 2–7). In PBC neurons, but not in NH neurons (b, 1–7), in addition to the NMDAR1-a splice variant, the NMDAR1-b splice variant (461 bp) is found very often (a, lanes 37–41). The C-terminal NMDAR1-1 splice variant (488 bp) is quite rare in all analyzed cell types (c, lanes 16–22; d, 8–14). NMDAR1-2 is expressed more strongly than NMDAR1-1 in the PBC (c, lanes 17–21) and in the NTS (d, lanes 8–12), but hardly detectable in the NH (data not shown). Expression of NMDAR1-3 (433 bp) was hardly detectable in PBC (e, lanes 36–42) and NH (f, lanes 8–14) neurons. In contrast, NMDAR1-4 (322 bp) was found to be much more abundant in all analyzed cell types (e, lanes 36–41; f, lanes 9–14). From total rat brain, in addition to PCR fragments for NMDAR1-3 and NMDAR1-4, a PCR product of intermediate size was obtained (e, f, rat brain) which is probably a PCR artefact as a result of hybridization of NMDAR1-3 and NMDAR1-4 fragments because of their striking similarity. Artifacts (e.g. c, cell 18 high molecular weight; d, cell 9 low molecular weight) were observed sometimes. To exclude false positive results because of these artifacts, all PCR fragments of the correct size were confirmed by restriction analysis.

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Table 4.  Statistical significance of differences between cell types in the detection of a given subunit
SubunitCell type 1Cell type 2SignificanceTest
  1. Cell type 1, subunit detected more often than in cell type 2; significance, statistical probability for the subunit being not more prominent in cell type 1 than in cell type 2; test 1, four arrays test; test 2, Fisher's exact test.

NMDAR1-bNHNTS< 0.0011
NMDAR1-2PBCNH< 0.011
NMDAR1-2NTSNH< 0.051
NR3APBCNTS< 0.0052
NR3APBCNH< 0.000052
NMDAR1-aPBCNH< 0.321
NMDAR1-bNHPBC< 0.241

In contrast to NMDAR1-a, for the NMDAR1-b splice variants significant differences between cell types were found (Table 4). NMDAR1-b splice variants were observed in 55 ± 5% of the PBC (Fig. 5a) and NH neurons, but only in 12% of the NTS neurons (Fig. 5b and Table 3).

Table 3.  Detection of NMDA receptor subunits in different brainstem cell types
Cell type SubunitPBCNHNTS
Detection rate (%)No. of cells analyzedDetection rate (%)No. of cells analyzedDetection rate (%)No. of cells analyzed

Less frequently than the expression of NMDAR1-a, expression of the C-terminal NMDAR1-1 splice variants was observed, but only in PBC neurons (Fig. 5c, cell 19) and not in the NH or the NTS (Fig. 5d).

Compared with NMDAR1-1 splice variants, NMDAR1-2 splice variants were much more abundant in the cell types analyzed. NMDAR1-2 splice variants were found in 45 ± 5% of the PBC (Fig. 5c) and NTS neurons (Fig. 5d), but only in 16% of the NH neurons. This difference is statistically significant (see Table 4).

Among all NMDAR1 splice variants, NMDAR1-3 variants turned out to be the least abundant being hardly detectable in the PBC and the NTS and undetectable in the NH (Figs 5e and f; Table 3).

Unlike NMDAR1-3 splice variants, NMDAR1-4 splice variants were found to be very prominent in all three cell types analyzed. NMDAR1-4 splice variants were detected in 60–75% of the PBC (Fig. 5e), the NH (Fig. 5f), and the NTS neurons.

Like NMDAR1-4 splice variants, NR2A is expressed in the PBC, the NH and the NTS neurons. With respect to NR2A, no differences were observed between all three cell types: NR2A expression was seen in 42 ± 3% of all those neurons (Table 3).

NR2B, like NR2A, turned out to be expressed in similar amounts in NH, NTS and PBC. NR2B was detected in 60–65% of the neurons (Table 3), thus much more often than NR2A. Examples for the detection of NR2B in PBC and NH neurons are shown in Figs 6(a and b), respectively.


Figure 6. Detection of NR2 and NR3A in single PBC and NH neurons. NR2B expression was observed in many PBC neurons (a, lanes 17–20) and NH neurons (b, lanes 29–35). No differences in the expression of NR2B showed up between the cell types. NR2D was detected in many PBC neurons (c, lanes 17–21) and NH neurons (d, lanes 29–35). In contrast to NR2D, differences in the expression of NR3A were observed between the cell types. NR3A was detectable in all PBC neurons (e, 36–42), but only in one-third of the NH (f, lanes 15–21) and NTS (data not shown) neurons. For other details see legend to Fig. 5.

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Among all NR2 subunits, the detection rate for NR2C was the lowest. In one-third of the PBC neurons NR2C was found, less often in the NH neurons, and barely in the NTS neurons (Table 3).

Like NR2A and NR2B, NR2D turned out to be expressed in similar amounts in all three cell types analyzed. NR2D expression was observed as often as expression of NR2B, in about 60% of the neurons (Figs 6c and d).

In contrast to NR2A, NR2B and NR2D, significant differences between the cell types was evident for expression of NR3A (Table 4). NR3A was detected in all analyzed PBC neurons (Fig. 6e), but only in one-third of the NH (Fig. 6f) and NTS neurons. The results are summarized in Tables 3–5.

Table 5.  NMDA receptor subunits of different brainstem cell types
  1. NMDA receptor subunits differing between NH and NTS are indicated in bold.

NMDAR1Subset of:  


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Comparison of present data with immunocytochemical, in situ hybridization and RT-PCR studies

The single cell PCR technique used in the present study allows the analysis of many different genes in parallel. Additionally, it has a high spatial resolution and thereby excludes false positive results as a result of the expression of glutamate receptor subunits in non-neuronal cells. In the present study, single neurons of the PBC, the NH and the NTS were analyzed for their NMDA receptor subunit composition. So far, no studies of the NMDA receptor subunit composition in identified single brainstem neurons have been published. Previous investigations concerning the expression of glutamate receptors in interneurons of the PBC neurons (Paarmann et al. 2000) neglected the differential expression of NMDAR1 splice variants and NR3A.

Various authors reported a strong expression of NMDAR1 in the PBC, the NH and the NTS in the brainstem of rat at the RNA level (Watanabe et al. 1994; Paarmann et al. 2000). The ratio between ‘a’ and ‘b’ splice variants of NMDAR1, however, has not yet been studied in brainstem, while in forebrain and cerebellum ratios of NMDAR1-a to NMDAR1-b of 5 : 1, and 1 : 5, respectively, were found (Anantharam et al. 1992; Sugihara et al. 1992). Laurie and Seeburg (1994) showed in an in situ hybridization study that both N-terminal splice variants are expressed in brainstem. However, the expression of these splice variants in identified brainstem nuclei or neurons was not investigated. The present study clearly demonstrates that NMDAR1-a splice variants are very prominent in all three cell types analyzed. In contrast, NMDAR1-b splice variants were only detected in the PBC and the NH, but hardly in the NTS (Table 3). Therefore, the detection of both NMDAR1-a and NMDAR1-b splice variants in brainstem neurons confirms the results of the in situ hybridization study of Laurie and Seeburg (1994).

The present study revealed that the C-terminal NMDAR1-1 splice variants are quite rare in brainstem neurons analyzed (Table 3). These results are consistent with the already-described weak expression of these splice variants in brainstem (Laurie and Seeburg 1994; Benke et al. 1995). In contrast, a strong expression of a different C-terminal splice variant, NMDAR1-2, has been previously demonstrated in brainstem (Laurie and Seeburg 1994), but not at the level of identified nuclei or neurons. So far, to study NMDAR1 expression in the PBC, the NH and the NTS, an antibody has been used which recognizes NMDAR1-1 as well as NMDAR1-2 splice variants (Petralia et al. 1994b; Ambalavanar et al. 1998). However, because NMDAR1-1 is quite rare in brainstem (Laurie and Seeburg 1994; Benke et al. 1995), the immunoreactivity detected by Petralia et al. (1994b) in rat and by Ambalavanar et al. (1998) in cat supposedly is almost exclusively because of the presence of NMDAR1-2. Ambalavanar et al. (1998) found a much weaker immunoreactivity in the NH compared with PBC and NTS. This finding fits exactly the results of the present single cell PCR study (Table 3). This study revealed that NMDAR1-2 splice variants are abundant in the PBC and NTS, but not in NH. In cat brainstem, however, with the same antibody used by Amabalavanar et al., in another immunocytochemical study no differences between the PBC, the NH and the NTS were observed (Petralia et al. 1994b), reflecting differences between organisms analyzed (cat vs. rat).

Much less prominent than NMDAR1-2 is a different C-terminal splice variant of NMDAR1, NMDAR1-3. So far, NMDAR1-3 expression has only been described in hippocampus and cortex, but not in brainstem. However, even in the hippocampus and the cortex, the other C-terminal splice variants are more prominent than NMDAR1-3 (Laurie and Seeburg 1994). This generally weak expression of NMDAR1-3 fits the very weak (PBC, NTS) or undetectable (NH) expression found in brainstem neurons in the present study (Table 3).

Conversely, NMDAR1-4 turned out to be very abundant in all three cell types analyzed in the present study (Table 3). This finding confirms the results of Laurie and Seeburg (1994) who described a strong expression of NMDAR1-4 in brainstem.

Compared with NMDAR1 splice variants, much more is known about NR2 subunit expression in PBC, NH and NTS. In mouse brainstem, expression of NR2A starts later than expression of NR2B. However, between P0 and P21, NR2B is largely replaced by NR2A (Watanabe et al. 1992). In P5 to P8 mice and in P21 mice, both NR2A and NR2B have been detected in PBC, NH and NTS, except for NR2B in PBC of P21 mice (Watanabe et al. 1994; Paarmann et al. 2000). These findings argue in favour of the immunoreactivity detected with an antibody specific for both NR2A and NR2B in all three nuclei in young rats (Petralia et al. 1994a) being caused by the presence of both subunits. Therefore, a strong expression of NR2A and NR2B demonstrated in the present study in P6 to P11 rats in all three cell types is similar to the findings of the previous studies.

In contrast to NR2A and NR2B, for NR2C slightly contradictory results have been reported so far. Watanabe et al. (1994) observed no expression of NR2C in these nuclei, whereas in a different study NR2C was detected in all three nuclei, although weaker in the PBC and NH (Paarmann et al. 2000). In the present study, which utilized P6-P11 rats, NR2C was found in PBC neurons at a similar level as described for P5 to P8 mice. However, differences between the two RT–PCR studies were apparent for the NTS. NR2C was reported to be strongly expressed in the NTS at the nuclei level (Paarmann et al. 2000), but was hardly detectable in single NTS neurons in the present study. These findings argue for a strong NR2C expression in non-neuronal cells in the NTS or for differences between P5 to P8 mice and P6 to P11 rats. In the NH, in single neurons a weak expression of NR2C was noted in this study, a finding that has been already described at the nuclei level (Paarmann et al. 2000).

The expression of NR2D in the brainstem has been previously demonstrated both at the RNA level (Watanabe et al. 1992; Ishii et al. 1993; Monyer et al. 1994), and at the protein level (Wenzel et al. 1995; Wenzel et al. 1996; Laurie et al. 1997). However, in an in situ hybridization study (Watanabe et al. 1994) detected no NR2D expression in the PBC, the NH and the NTS of P21 mice. In contrast, in a different study strong expression of NR2D was seen at the level of the nuclei in the PBC, the NH and the NTS, and even in single neurons of the PBC (Paarmann et al. 2000). In the present study NR2D turned out to be very abundant in all three analyzed cell types, a finding which is in line with this previous report. Therefore, the observed differences between the two RT–PCR studies and the in situ hybridization study might be explained by a higher sensitivity of RT–PCR, and a stronger NR2D expression as a result of a different developmental stage being analyzed (Monyer et al. 1994).

In contrast to the NR2 subunits, NR3A expression differed markedly between the cell types studied here. NR3A was found to be much more abundant in PBC neurons compared with NH and NTS neurons (Table 3). The expression of NR3A in the PBC, NH and NTS has been previously reported (Paarmann et al. 2000) and was further confirmed in the present study. As expression of NR3A is strongly dependent on the developmental stage (Sucher et al. 1995), the much stronger expression of NR3A in PBC neurons compared with NH and NTS neurons, which was not observed in a different RT–PCR study (Paarmann et al. 2000), is probably caused by differences in the developmental state of the experimental animals (6- to 11-day-old rats in this study compared with 5- to 8-day-old mice in the previous study).

Many previously analyzed neurons show relatively slow kinetics of the NMDA currents (Table 2). In comparison with the fast decay times found for hypoglossal neurons, which have also been described by other groups (O'Brien et al. 1997), and PBC neurons, decay times in the NTS were approximately 10 times slower. In some cases, the kinetics of NMDA currents could only be fitted adequately using two exponential functions, indicating a biphasic current profile.

Functional implications

The expression of different C-terminal NMDAR1 splice variants may affect binding of NMDAR1 to intracellular proteins. The C1-cassette, present in NMDAR1-1 and NMDAR1-3, has been reported to bind several proteins such as neurofilaments (Ehlers et al. 1998), calmodulin (Ehlers et al. 1996) and yotiao (Lin et al. 1998). Furthermore, it contains an ER retention signal (Standley et al. 2000) and a nuclear localization signal (Holmes et al. 2002). Ca2+-activated calmodulin can reduce the mean open time of NMDA receptors containing NMDAR1-1 and NMDAR1-3 (Ehlers et al. 1996). Therefore, as the C1 cassette was hardly detectable in this study, the NMDA receptors of PBC, NH and NH are probably largely resistant to this direct inhibitory effect of calmodulin and, because of the lack of the ER retention signal, can be easily transported to the cell surface. Additionally, the putative proteolytic cleavage and redirection of the C-terminal domain to the nucleus is very unlikely to occur in the NH and the NTS. In contrast to the C1-cassette, the function of the C2-cassette is not yet clear. NMDAR1-2 and NMDAR1-4 do not only differ in the presence or absence, respectively, of the C2-cassette, but also in their C-termini (Sugihara et al. 1992; Hollmann et al. 1993). Although literature about a direct interaction between the C-terminus of NMDAR1-4 and a PDZ domain of PSD95 is contradictory (Kornau et al. 1995; Bassand et al. 1999), the C-terminus is able to bind COPII (Mu et al. 2003) and is likely capable of binding to a PDZ domain protein (Standley et al. 2000). The presence of the COPII binding site in NMDAR1-4 further facilitates the transport of the NMDA receptors of the PBC, the NH, and the NTS to the cell surface. Because of the much stronger expression of NMDAR1-b splice variants in the PBC and the NH, their NMDA receptors might be regulated in a different manner compared with neurons showing slower deactivation time courses, like those in the NTS. The NMDA receptors of NH and PBC are supposedly more susceptible to protein kinase C-mediated potentiation (Durand et al. 1993), but less to potentiation by zinc (Hollmann et al. 1993), neurosteroids (Malayev et al. 1998; Ceccon et al. 2001) and polyamines (Zheng et al. 1994).

The present study clearly demonstrates that all three analyzed cell types have distinct expression patterns regarding their NMDAR1 splice variants, but not their NR2 subunits. Because NH and NTS cells differ only in the expression of one NMDAR1 splice variant, NMDAR1-4b in NH compared with NMDAR1-2a in NTS (Table 5), this difference might provide a possible molecular basis for the observed differences in the decay time of deactivation of NMDA EPSCs between NTS [+40 mV, 300 ms (Titz and Keller 1997)] and the much faster NH [+40 mV, 126 ms (O'Brien et al. 1997)]. Decay times of deactivation of NMDA receptors have been previously studied in recombinant expression systems (Monyer et al. 1992, 1994; Vicini et al. 1998; Rumbaugh et al. 2000). They have been shown to depend on both the NR2 subunits as well as on the NR1 splice variant. Co-expression of the five most prominent NMDAR1 splice variants with NR2A did not reveal any significant differences (Vicini et al. 1998). However, upon co-expression with NR2B, the presence of the NMDAR1-b splice variants leads to a faster deactivation compared with the corresponding NMDAR1-a splice variants (Rumbaugh et al. 2000). Our single cell PCR data are consistent with a simple model featuring a tetrameric NMDA receptor consisting of one NR2B, one NR2D, one NMDAR1-4a, and one additional NMDAR1 subunit depending on the specific cell type. In the NTS, this additional subunit would be NMDAR1-2a, in the NH NMDAR1-4b, and in the PBC, which like NH shows very fast decay times of deactivation (Table 2), NMDAR1-2b. However, in the PBC different NMDAR1 splice variants such as NMDAR1-4b and NMDAR1-2a cannot be excluded, because C- and N-terminal splice variants were analyzed independently. Additionally, this model does not take into account the strong NR3A expression in the PBC. Using co-immunoprecipitation of NR3A with NMDAR1 and NR2B, Das et al. (1998) showed that NR3A is in fact part of a functional NMDA receptor complex. However, so far no studies have been published which determined the ratio between NR3A and NMDAR1 and NR2 in a functional NMDA receptor complex.

Taken together, our data suggest that the fast kinetics of the NMDA EPSCs in the PBC and the NH compared with the NTS could be achieved by a much stronger expression of NMDAR1-b splice variants. Thus, the decay times of deactivation of the NMDA receptors might not, as previously believed, depend solely on the NR2 subunits, but also on the NMDAR1 splice variants.

The present study clearly showed that the frequency of breathing in young rats can exceed 5 Hz. Therefore, to fulfil a modulatory function in each cycle of breathing, NMDA receptors have to act within 200 ms. Out of all analyzed different types of neurons, only the NMDA receptors expressed in the PBC and the NH, as demonstrated in this study, are able to act within this short timescale, which might be achieved by expression of unique combinations of NMDA receptor subunits. The results presented here delineate the NMDA receptor subunits and splice variants expressed in respiratory-related and control cells; however, the assembly of the splice variants and the spatial distribution of receptors remain unknown. In addition to the differential pattern of NMDA receptor mRNA expression, other processes such as post-translational modification could potentially modulate the specific properties of NMDA receptors. Our conclusion from the data presented is that the observed differential expression of one NMDA receptor subunit might regulate fast kinetics of synaptic currents.


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

We thank D. Richter for valuable discussions and support. This research was partially supported by DFG grants to the SFB 406 and the CMPB Göttingen.


  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
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