Intrinsic Optical Signals in Respiratory Brain Stem Regions of Mice: Neurotransmitters, Neuromodulators, and Metabolic Stress

M. Haller, S. L. Mironov, and D. W. Richter

Physiologisches Institut, Georg-August-Universität Göttingen, D-37073 Gottingen, Germany


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INTRODUCTION
METHODS
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DISCUSSION
REFERENCES

Haller, M., S. L. Mironov, and D. W. Richter. Intrinsic Optical Signals in Respiratory Brain Stem Regions of Mice: Neurotransmitters, Neuromodulators, and Metabolic Stress. J. Neurophysiol. 86: 412-421, 2001. In the rhythmic brain stem slice preparation, spontaneous respiratory activity is generated endogenously and can be recorded as output activity from hypoglossal XII rootlets. Here we combine these recordings with measurements of the intrinsic optical signal (IOS) of cells in the regions of the periambigual region and nucleus hypoglossus of the rhythmic slice preparation. The IOS, which reflects changes of infrared light transmittance and scattering, has been previously employed as an indirect sensor for activity-related changes in cell metabolism. The IOS is believed to be primarily caused by cell volume changes, but it has also been associated with other morphological changes such as dendritic beading during prolonged neuronal excitation or mitochondrial swelling. An increase of the extracellular K+ concentration from 3 to 9 mM, as well as superfusion with hypotonic solution induced a marked increase of the IOS, whereas a decrease in extracellular K+ or superfusion with hypertonic solution had the opposite effect. During tissue anoxia, elicited by superfusion of N2-gassed solution, the biphasic response of the respiratory activity was accompanied by a continuous rise in the IOS. On reoxygenation, the IOS returned to control levels. Cells located at the surface of the slice were observed to swell during periods of anoxia. The region of the nucleus hypoglossus exhibited faster and larger IOS changes than the periambigual region, which presumably reflects differences in sensitivities of these neurons to metabolic stress. To analyze the components of the hypoxic IOS response, we investigated the IOS after application of neurotransmitters known to be released in increasing amounts during hypoxia. Indeed, glutamate application induced an IOS increase, whereas adenosine slightly reduced the IOS. The IOS response to hypoxia was diminished after application of glutamate uptake blockers, indicating that glutamate contributes to the hypoxic IOS. Blockade of the Na+/K+-ATPase by ouabain did not provoke a hypoxia-like IOS change. The influences of KATP channels were analyzed, because they contribute significantly to the modulation of neuronal excitability during hypoxia. IOS responses obtained during manipulation of KATP channel activity could be explained only by implicating mitochondrial volume changes mediated by mitochondrial KATP channels. In conclusion, the hypoxic IOS response can be interpreted as a result of cell and mitochondrial swelling. Cell swelling can be attributed to hypoxic release of neurotransmitters and neuromodulators and to inhibition of Na+/K+-pump activity.


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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The intrinsic optical signal (IOS) provides a noninvasive technique for imaging "neuronal functions" in living brain. The signal originates from changes in the refractory indexes of the cytoplasm and the extracellular space that together determine light transmittance and/or scattering. As the signal originates from endogenous processes in the tissue, it does not require any cell labeling and is thus not subject to bleaching. The IOS imaging technique has been applied to the intact brain as well as to slice preparations. In the intact brain, IOS has been used, e.g., to map the spatial distribution and patterning of neuronal activity (Holthoff and Witte 1998) as well as to verify propagation of seizure (Federico et al. 1994). In slices from hippocampus, neocortex, and the retina, the IOS technique was used to examine various pathological processes, such as spreading depression in the retina (Fernandes de Lima et al. 1997; Ulmer et al. 1995) and the hippocampus (Muller and Somjen 1998; Obeidat and Andrew 1998) or excitotoxicity originating from enhanced release of excitatory neurotransmitters (Andrew et al. 1999).

The mechanisms underlying the IOS are complex, and its origin is as yet not completely understood. There are strong indications that an essential component of IOS is determined by changes in cell volume (Andrew et al. 1996; Polischuk and Andrew 1996), which might originate from cell swelling due to Na+, Ca2+, Cl- accumulation and concomitant water fluxes through the plasma membranes (Andrew and MacVicar 1994) or from swelling of adjacent glial cells buffering extracellular K+ that is released during neuronal activity (MacVicar and Hochman 1991). However, there is also evidence that not all components of the IOS can be explained in terms of cell volume changes. Buchheim et al. (1999) found that electrical hyperactivity in hippocampal slices of rat leads to diverse changes in the IOS: the IOS also increased during shrinkage of the extracellular space, indicating the presence of a volume-independent mechanism of IOS changes. Similarly, in hippocampal slices (Aitken et al. 1999), light transmittance was found to increase during moderate hypotonia, but to decrease during pronounced arterial hypotonia even though cells continued to swell. Another puzzling information is that spreading depression (induced, for example, by potassium injection) and hypoxic spreading depression-like depolarization (induced by oxygen withdrawal) lead to a decrease in transmitted light intensity (Muller and Somjen 1999).

A possible explanation for all these divergent phenomena could be that in addition to cell volume changes, IOS is also affected by changes in the distribution and/or volume of subcellular structures. Objects such as dendritic spines, dendritic beads and cellular organelles may change their shape as water enters or leaves the cell. Specifically beading of the dendritic arbor, i.e., the formation of varicosities in dendrites as a result of an excitotoxic insult (Andrew et al. 1999; Hasbani et al. 1998; Polischuk et al. 1998), and the swelling of mitochondria (Aitken et al. 1999; Andrew et al. 1999) have been correlated with increase in light scattering.

In the present study, IOS was used to measure light transmittance through a brain stem slice preparation that contains a functional network generating ongoing rhythmic activity (Smith et al. 1991). We aimed to monitor changes within distinct regions containing functional networks such as the nucleus hypoglossus and the periambigual region, which is involved in respiratory rhythm generation and pattern formation. We investigated changes of the IOS during normoxia, but also during hypoxia and disturbances of osmotic pressure. Specifically, we focused on the mechanisms that contribute to the hypoxic IOS-response such as glutamate release and blockade of the Na+, K+ pump. Finally, we examined the effects of drugs, acting on KATP channels. These channels are modulated by intracellular ATP levels (Haller et al. 1999) and thus directly couple cellular metabolism to modulation of membrane permeability and neuronal excitability. KATP channels are believed to modulate the respiratory activity (Pierrefiche et al. 1996) and to play a vital role in the mechanisms protecting against hypoxic cell damage (Krishnan et al. 1995; Mironov et al. 1998; Murphy and Greenfield 1991). Our data indicate that neuronal activity together with activity-dependent ion transportation and presumably mitochondrial swelling contribute essentially to the IOS.


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Slice preparations

Experiments were performed on the medullary slice preparation from neonatal mouse (Mironov et al. 1998; Smith et al. 1991). The preparation contains a functional respiratory network that generates ongoing rhythmic activity. All animals were housed, cared for, and killed in accordance to the recommendations of the European Commission (No. L 358, ISSN 0378-6978), and protocols were approved by the Committee on Animal Research, Göttingen University. The brain stem-spinal cord was isolated in ice-cold artificial cerebrospinal fluid (ACSF, composition listed below), and a single transverse 700-µm-thick slice containing the pre-Bötzinger complex was cut from the brain stem, transferred to the recording chamber, and mounted on the stage of an upright microscope (Axioscop FS, Zeiss). The slice was fully submerged in a continuously flowing ACSF (28°C, 40-50 ml/min) and gassed with carbogen (95% O2-5% CO2). To prevent diffusional loss of dissolved gases, the perfusing solution was delivered to the experimental chamber via stainless steel tubing. The respiratory rhythm was established at an elevated extracellular K+ concentration of 8 mM. One hypoglossal (XII) rootlet was sucked into a blunt capillary for extracellular recording of rhythmic respiratory discharges. XII activity was amplified 5,000-10,000 times, band-pass (0.25-1.5 kHz) filtered, rectified, and integrated (Paynter filter with a time constant of 50-100 ms). This integrated version of nerve activity typically has an amplitude of 0.1 mV. However, as the amplitude is quite variable and depends on the leak conductance of the suction electrode, a relative calibration was made assuming a control peak amplitude of 1 to describe temporal changes of nerve activity during individual experiments.

All drugs were added directly to the bath reservoir and arrived at the experimental chamber after a delay of 8-12 s. Drug wash out was achieved by perfusing 400-500 ml fresh solution containing the same [K+]o. To induce hypoxic conditions in the slice, the bubbling gas mixture was changed from carbogen to 95% N2 and 5% CO2. The oxygen level was measured with oxygen-sensitive electrodes (Diamond Electro-Tech, Ann Arbor, MI) as described previously (Mironov et al. 1998). PO2 electrodes were placed 100 ± 25 (SE) µm below the slice surface, into a region where inspiratory neurons were located. Fifteen to 20 s after exchanging oxygen in the perfusing solution by nitrogen, extracellular PO2 changed from 232 ± 39 to 6 ± 4 mmHg and then remained constant (n = 15, P < 0.05).

Imaging techniques

For IOS measurements, slices were transiently transilluminated using a tungsten lamp that was controlled by a voltage-regulated power supply (Zeiss). Near-infrared illumination was obtained using a highpass filter, which cut off light below 780 nm wavelength. Water immersion objectives were used to avoid light scattering at the air/tissue interface, which otherwise would have distorted the IOS significantly (Kreisman et al. 1995). Objective lenses with different magnifications (×2.5, ×10, ×63) of an upright microscope were used to obtain images of the whole slice or single cells. Images were collected with a charge-coupled device (CCD) camera (Princeton Instruments) at rates of up to 1 frame/s. In the experiments, we followed the procedure described by Andrew and MacVicar (1994). First, control images were recorded, areas of interest were defined, and averages of the transmitted light were used as control values, To. The data were transformed into relative changes, according to IOS = Delta T/To. Images were presented using a pseudocolor intensity scale.

For measuring single-cell volume (Fig. 1C), inspiratory neurons, i.e., neurons that discharged rhythmic bursts of action potentials in synchrony with hypoglossal (XII) nerve activity, were filled with mag-fura-2 via the patch pipettes and illuminated with a monochromator (Till Photonics, Planegg, Germany) with the ultraviolet (UV) light of 346-nm wavelength, which corresponds to the isosbestic point of mag-fura-2 (Raju et al. 1989).



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Fig. 1. Intrinsic optical signal and cell volume changes during hypoxia. A: subtraction image of slice. Use of the subtraction algorithm eliminates the nylon mesh and the stabilizing "horse-shoe" from the infrared image. The boxes mark the regions of the nucleus hypoglossus and periambigual region displayed in pseudocolor in B. B: the intrinsic optical signal (IOS) displayed a reversible hypoxic increase of up to 40% (n = 11). C: during hypoxia an inspiratory neuron filled with mag-fura-2 was observed to swell (n = 3). The images on the left display pictures take before (top) and during hypoxia (bottom), which were then subjected to a threshold procedure (right). Finally, the control picture is subtracted from the hypoxia picture. The resulting image, which clearly demonstrates a volume increase, is displayed in pseudocolor below.

Solution and drugs

ACSF contained (in mM) 128 NaCl, 3 KCl, 1.5 CaCl2, 1.0 MgSO4, 21 NaHCO3, 0.5 NaH2PO4, and 30 D-glucose, pH adjusted to 7.4 with NaOH. Solutions with variable K+-concentration were obtained by replacing NaCl with KCl. Intracellular solution used for whole cell recording contained (in mM) 120 K+-gluconate, 15 NaCl, 2 MgCl2, 10 HEPES, 0.5 Na2ATP, 1 CaCl2, 3 1,2-bis-(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), and sometimes 100 µM mag-fura-2. The pH was adjusted to 7.4 with KOH. Solution osmolarity ranged from 285 to 290 mosM. HMR1098 was kindly provided by Aventis Pharma Deutschland GmbH (Frankfurt, Germany). All other reagents were obtained from Sigma-Aldrich (Deisenhofen, Germany).


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IOS during cell swelling

Figure 1A shows the infrared image of the preparation at low magnification. IOS measurements focused on two regions that reveal rhythmic respiratory activity, i.e., the periambigual region (pa) and the nucleus hypoglossus (nh) as defined by rectangles in Fig. 1A. The relative changes in IOS in these regions during hypoxia are displayed in Fig. 1B using a pseudocolor scale (n = 11). In both regions, the IOS revealed a pronounced increase within 4 min after oxygen depletion and recovered completely after 15 min of reoxygenation.

Individual cells were observed to swell during hypoxia (Fig. 1C). Cell volume changes were monitored in identified inspiratory neurons loaded with mag-fura-2 (n = 3). The dye was excited at its isosbestic point for Ca2+ binding (346 nm) to eliminate the contribution of the hypoxic rise in Ca2+ to the fluorescence signal. The cell contours were defined from corresponding cell masks obtained by setting a threshold light intensity filter (right-hand side of Fig. 1C). The bottom panel represents the difference between the mask obtained after 4 min of hypoxia and that of control. Thus it indicates the area of volume increase of the cell as highlighted by pink (positive) color.

The IOS changed during various maneuvers that induced changes in respiratory activity. Figure 2 shows simultaneous measurements of the respiratory output as indicated in hypoglossal nerve activity (XII) and the corresponding IOS obtained from the nucleus hypoglossus and the periambigual region (gray traces). Increase of K+ concentration within the external bath solution from 3 to 9 mM (Fig. 2A) resulted in the appearance of rhythmic respiratory activity, which was accompanied by a significant rise of IOS. The IOS changes were significantly larger in the nucleus hypoglossus as compared with the changes in the periambigual region indicating a differential effect of ongoing neuronal and metabolic activity. On re-superfusion with low [K+] solution (3 mM), IOS values returned to baseline levels. The IOS was also reversibly changed during variations in osmolarity of the perfusion solution (Fig. 2B). In slices exposed to a hyperosmotic solution (+15 mosM) obtained by supplementation of mannitol, the IOS declined, whereas there was a rise in IOS in hyposmotic medium (-15 mosM). In both cases the changes of IOS were comparable in size in both regions and returned to control levels within 15 min after restoration of isosmotic conditions. Thus as expected, osmotic cell swelling caused an increase in light transmittance, whereas shrinkage resulted in a decrease.



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Fig. 2. Hypoglossal nerve activity and IOS recorded from nucleus hypoglossus (nh) and periambigual region (pa). A: when extracellular K+ was elevated the respiratory rhythm appeared (top trace) and the IOS (bottom trace) increased both in the periambigual region (pa) and the nucleus hypoglossus (nh; n = 6). B: the IOS rose during hyposmotic conditions and fell during hyperosmotic conditions (n = 4). C: during hypoxia, the IOS increased and returned to control levels on reoxygenation (n = 18). The same profile was displayed during chemical hypoxia (n = 2) with a perfusion time of 14 min for KCN (D). The respiratory rhythm (XII) exhibits a biphasic response to hypoxia consisting of an initial augmentation and a subsequent depression (C and D, top traces). The derivative of Delta T/To (C and D, bottom traces) shows that the fastest change in IOS occurs approximately toward the end of the augmentation phase of hypoxia and that rate of increase in IOS is greater for periambigual region (pa) than for nucleus hypoglossus (nh). A relative calibration is used to describe the temporal changes of hypoglossal nerve activity (XII) allocating a peak amplitude of 1 to control conditions.

Hypoxic IOS response

Hypoxia leads to an initial augmentation of respiratory activity followed by a secondary depression superimposed on a slowly developing DC signal (Cherniack et al. 1970; Richter et al. 1991) (Fig. 2C). The IOS rose during both phases of hypoxia even though the rate of IOS rise decreased with time. In some cases the IOS leveled off. As previously reported (Volker et al. 1995) metabolic poisoning with cyanide elicits responses similar to hypoxia (Fig. 2D) and occludes subsequent hypoxic reactions. Respiratory activity and IOS, however, recovered fully after reoxygenation or wash out of KCN.

There were important differences in the IOS responses obtained from the two brain stem regions. During the initial phase of hypoxia and after changing extracellular K+, the latency was shorter, and the amplitude as well as the rate of IOS rise were larger in the nucleus hypoglossus as compared with the periambigual region (Fig. 2, A, C, and D). However, there were no such regional differences after changing the osmolarity of the perfusion solution (Fig. 2B).

Hypoxia induces augmented release of excitatory and inhibitory neurotransmitters and neuromodulators (Richter et al. 1999), which might contribute to the observed hypoxic IOS responses. The principal excitatory neurotransmitter involved in the early phase of hypoxia is glutamate, which activates N-methyl-D-aspartate (NMDA) and kainate/alpha -amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptor subtypes (Choi 1993; Schurr et al. 1995). Thus to simulate initial hypoxic responses, NMDA and kainate were applied to the slice preparation, which resulted in a marked, but reversible increase in IOS similar to the hypoxic IOS response. The effects were blocked by the specific antagonists 2-amino-5-phosphonovaleate (AP-5) and 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX; Fig. 3, A and B). Glutamate application led to a transient rise of the IOS (Fig. 3C). Adenosine, which is released during hypoxic depression (Richter et al. 1999), induced a transient decrease of the IOS (Fig. 3D).



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Fig. 3. Effect of application of neurotransmitters on IOS. Application of 50 µM N-methyl-D-aspartate (NMDA; A) and 50 µM kainate (B) led to an increase in IOS, which could be blocked by 50 µM 2-amino-5-phosphonovaleate (AP-5) and 3.5 µM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), respectively (n = 2). C: high concentrations of glutamate (500 µM, n = 3) resulted in transient IOS rises. D: 200 µM adenosine led to a small transient decrease in IOS (n = 3). E: after application of glutamate uptake blocker L-trans-pyrrolidine-2,4-dicarboxylic acid (PDC; 30 µM, n = 3) the IOS decreased reversibly (top). Successive periods of hypoxia displayed a pronounced reversible decrease in hypoxic IOS response amplitude as becomes evident when the baseline was subtracted (bottom).

When the glutamate uptake blocker L-trans-pyrrolidine-2,4-dicarboxylic acid (PDC) was added, the IOS baseline level decreased, and consecutive periods of anoxia elicited only diminished IOS responses (Fig. 3E). On wash out, the IOS returned to control levels, and the hypoxic IOS response recovered. To further investigate the correlation between IOS and neurotransmitter release, we applied bafilomycin A1, a potent and specific blocker of the vacuolar-type (V-type) ATPase, which eliminates the driving force for the uptake of glutamate, serotonin, and GABA into synaptic vesicles (Moriyama and Futai 1990; Zhou et al. 2000). Again we observed a reversible decrease of the IOS as well as a partial blockade of the hypoxic IOS response (Fig. 4A).



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Fig. 4. Effect of bafilomycin, ouabain, furosemide, and tetrodotoxin (TTX) on IOS. A: the glutamate uptake into synaptic vesicles was reduced by applying 50 nM bafilomycin A1, a blocker of vacuolar-type (V-type) ATPase. Following bafilomycin A1 application the IOS decreased and the IOS amplitude of successive episodes of hypoxia was diminished. On wash out the IOS returned to control levels (n = 3). B: changes to the respiratory rhythm and the IOS after application of 50 µM ouabain. The IOS exhibited a slow decrease, and the hypoxic IOS response was augmented (n = 5, 50-100 µM ouabain). The respiratory rhythm displayed a hypoxia-like response to ouabain application and was subsequently blocked. C: when Ca2+ channels were blocked with Cd2+, ouabain elicited a transient IOS increase (n = 2). D: after furosemide application (n = 3) the IOS exhibited a slow decrease. The hypoxic IOS response was not affected. E: following application of 1 µM TTX, the IOS decreased slowly. Hypoxic responses were partially blocked (n = 3).

Another source of the hypoxic IOS increases might be cell swelling due to ATP depletion and depression of Na+/K+-ATPase activity in the plasma membrane, which leads to accumulation of intracellular Na+ and loss of intracellular K+. To test this assumption the Na+/K+-ATPase blocker ouabain was applied. On ouabain application (50-100 µM), the respiratory activity displayed a hypoxia-like reaction, but the IOS baseline decreased (Fig. 4B). In the presence of ouabain, hypoxic stimuli elicited only 1-2 sequential IOS responses until the reaction of both the respiratory output and IOS were completely suppressed and did not recover even after 1 h of wash out. The final IOS response exhibited a higher amplitude and a delayed recovery on reoxygenation. The ouabain-induced decrease of IOS could be caused by cell shrinkage due to Ca2+ entry and subsequent activation Ca2+-activated K+ channels (Alvarez-Leefmans et al. 1994; Smith et al. 1993). To test this hypothesis, Ca2+ channels were blocked with Cd2+ before ouabain application. Indeed this resulted in a pronounced rise in IOS, which was transient and followed by a prolonged IOS decrease (Fig. 4C).

An important mechanism of volume regulation in glial cells is the Na+/K+/2Cl- cotransporter, which mediates the electroneutral uptake of ions (Haas 1994; Kimelberg et al. 1986) and thus contributes to cell volume regulation (Baba 1992; Chen et al. 1992; Hoffmann 1992; Walz 1992). Blockade of the Na+/K+/2Cl- cotransporter by furosemide should lead to fall of intracellular Na+, K+ and Cl- concentrations followed by water efflux and thus shrinkage of glial cells. Indeed we observed a decrease in the IOS after furosemide application, which was reversible after 60-80 min wash out. The hypoxic responses of IOS and respiratory activity remained unmodified (Fig. 4D), indicating that the Na+/K+/2Cl- cotransporter does not directly affect hypoxic responses of neuronal activity.

Blocking neuronal activity with tetrodotoxin (TTX) completely suppressed respiratory activity and slowly decreased IOS (Fig. 4E). The hypoxic IOS responses became progressively smaller, but they were not completely suppressed even after cessation of rhythmic output activity.

IOS response following application of KATP channel drugs

Normoxic and hypoxic respiratory activities are effectively modulated by KATP channels (Mironov et al. 1998; Pierrefiche et al. 1996). Therefore we tested the contribution of KATP channels to the IOS responses by applying channel-specific drugs, such as glibenclamide, tolbutamide (blockers), diazoxide, and pinacidil (openers) under normoxic conditions. High concentrations of such KATP channel-directed drugs affected the size of integrated XII bursts as exemplified in Fig. 5A. Application of the channel opener diazoxide (300-500 µM, n = 7) led to a decrease of the area of integrated hypoglossal burst activity by 21.2 ± 10.3%, whereas tolbutamide (300-500 µM, n = 7) and glibenclamide (30-50 µM, n = 7) resulted in an increase by 15.9 ± 12.8% and 6.4 ± 6.2%, respectively (Student's t-test, P < 0.05). We expected that the IOS would rise on application of KATP channel blockers due to enhancement of neuronal activity and cell swelling, and that it would correspondingly decline on application of KATP channel activators. The results, however, were variable. Both the applications of the agonist diazoxide (n = 28) and the blocker glibenclamide (n = 23) resulted in an increase in IOS in approximately one-half of the tests, but in a decrease in the remainder trials. Individual responses seemed to be specific for a given slice preparation in that they could be reproduced over a time span of several hours. These findings indicate that the mechanisms underlying generation of the IOS involve also processes other than neuronal or glial swelling.



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Fig. 5. KATP channel drugs and IOS. A: response of integrated hypoglossal nerve (XII) activity to application of KATP channel drugs diazoxide and glibenclamide. B: following 10 min of hypoxia, the application of KATP channel drugs led to an increase in IOS after diazoxide and a decrease after glibenclamide (n = 4, different preparations). C: application of plasmalemmal KATP channel blocker HMR 1098 leads to a reversible increase in IOS both during normoxia (n = 5) and hypoxia (n = 3). D: application of the mitochondrial uncoupler carbonylcyanide m-chlorophenylhydrazone (CCCP) resulted in a transient increase in IOS (n = 3).

One additional process might be mitochondrial swelling as KATP channels expressed in mitochondrial membranes (Hu et al. 1999; Wang and Ashraf 1999) are also affected by diazoxide and glibenclamide. To test this possibility, comparable metabolic conditions were established by 10 min of hypoxia followed by the application of KATP channel-directed drugs (Fig. 5B). In this case, the IOS started to rise during hypoxia and then consistently rose further when the channel opener diazoxide was applied or decreased after application of the channel blocker glibenclamide in all tests performed (n = 8). When HMR1098, which exclusively blocks plasmalemmal KATP channels, was applied (Fig. 5C), a reversible increase in IOS was observed in all slices both during normoxia (n = 5) and hypoxia (n = 3). The involvement of mitochondria was also verified by applying a mitochondrial uncoupler, carbonylcyanide m-chlorophenylhydrazone, CCCP. Figure 5D illustrates how CCCP application led to an initial increase in IOS, which then turned into a decrease toward levels well below control (n = 3). The latter effect manifested severe damage of cells, as the respiratory output was irreversibly lost.


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We used the IOS to monitor metabolic changes in neurons during hypoxia, the application of glutamate receptor agonists and KATP channel blockers or openers. The measurements were complicated by the fact that processes other than cell volume changes appeared to contribute significantly to IOS, necessitating a more detailed investigation of the origin of IOS in the rhythmic slice preparation.

Changes in cell volume

It is commonly accepted that any change in intracellular and/or extracellular osmolarity induce changes in cell volume (Andrew and MacVicar 1994; Lipton 1973). In this work, changes in osmolarity also modulated the IOS in the rhythmic brain stem slice preparation, verifying that part of the IOS reflects cell volume changes. This finding was substantiated by analyses of identified inspiratory neurons, which exhibited a pronounced and reversible increase in volume in parallel with the IOS rise during hypoxia. The effect was similar to that described for hippocampal neurons in slices (Kreisman and LaManna 1999; Turner et al. 1995).

Regional differences in IOS

There are significant differences between neuronal systems indicating differences in metabolic demands and/or protective processes. The measurements of faster and larger IOS changes in hypoglossal nucleus are consistent with the observations in previous studies (Haddad and Donnelly 1990; O'Reilly et al. 1995; Pierrefiche et al. 1997), that hypoglossal motoneurons have a low tolerance to hypoxia. In contrast to hippocampal neurons and other brain stem neurons, such as respiratory or dorsal vagal neurons, hypoglossal neurons exhibited a pronounced anoxic depolarization in response to O2 deprivation (Donnelly et al. 1992; O'Reilly et al. 1995; Pierrefiche et al. 1997; Richter et al. 1991, 1992). A larger anoxic depolarization in hypoglossal neurons would be expected to lead to a higher degree of cell swelling due to a larger influx of ions. Indeed, comparison of respiratory neurons with hypoglossal motoneurons revealed pertinent differences in the hypoxic IOS responses. After changes in bath osmolarity (±15 mosM), however, the IOS displayed no regional diversity, which indicates that passive water movement imposed by osmolarity changes affects the volume of all cell populations in a similar manner.

Mechanisms underlying the hypoxic IOS-response

As mentioned above, hypoxia has been observed to elicit a pronounced IOS response that is in part due to cell swelling. The most likely mechanisms underlying hypoxic cell swelling are Na+, Ca2+, and Cl- influxes (Haddad and Donnelly 1990; Mercuri et al. 1994) and failure of the Na+/K+ pump following ATP depletion (Calabresi et al. 1995; Le Corronc et al. 1999). After reoxygenation, the IOS recovered, indicating the restoration of the original cell volume. Important factors contributing to such regulatory volume decrease are Na+/K+ pump activity (Basavappa et al. 1998; Fraser and Swanson 1994) or parallel operation of Cl- and K+ channels, allowing efflux of K+ and Cl- (Hoffmann 1992; Lippmann et al. 1995). The various factors causing cell swelling and thus contributing to the hypoxic IOS increase were investigated by application of neurotransmitters, neurotransmitter uptake blockers, and blockade of Na+/K+ pumping.

EFFECT OF NEUROTRANSMITTERS ON IOS. Our experiments verified that processes that enhance neuronal activity, such as glutamate receptor activation, produce a transient increase in IOS, whereas adenosine (an agent that depresses spontaneous neuronal activity) transiently decreases IOS. The transient nature of the effect can be explained by immediate neurotransmitter reuptake. During hypoxia a prolonged accumulation of neurotransmitters and neuromodulation is expected due to impairment of uptake processes and catabolic enzymes in neurons and glial cells (Haddad and Jiang 1993; Neubauer et al. 1990). During the early hypoxic augmentation phase, predominantly glutamate and GABA are released, which is followed by a delayed release of adenosine and serotonin during hypoxic respiratory depression (Richter et al. 1999). Consequently, we assume that a glutamate-induced IOS increase contributes to the early hypoxic rise in IOS, whereas an adenosine-mediated fall contributes to the decreasing slope of the hypoxic IOS change that was observed during the depressive phase of hypoxia. This assumption is corroborated by the finding that a reduction in glutamate release, either induced by the glutamate uptake blocker PDC (Fig. 3E) or vacuolar ATPase inhibitor bafilomycin A1 (Fig. 4A), resulted in a general decline of IOS and a partial blockade of the hypoxic IOS response.

However, glutamate release leading to Na+ and Ca2+ fluxes into neurons and thus water flow across the membrane is probably not the only reason for IOS changes during hypoxia. We assume that other metabolic factors and structural changes of intracellular organelles also play an important role.

NA+/K+-PUMP AND NEURONAL ACTIVITY. It is generally accepted that depression of Na+/K+ pump activity leads to cell swelling due to accumulation of intracellular Na+ (Buckley et al. 1999; Shimizu and Nakamura 1992), thereby causing an increase in IOS similar to that observed during hypoxia. Surprisingly, blocking of Na+/K+ pump activity with ouabain did not mimic the hypoxic IOS response, but rather markedly decreased IOS baseline levels. This is consistent with the observations in a number of studies (Alvarez-Leefmans et al. 1994; Smith et al. 1993) that ouabain induces cell shrinkage rather than cell swelling. The authors suggested that ouabain induces a transient elevation of [Ca2+]i, which in turn activates a K+ efflux through Ca2+-activated K+ channels, leading to water loss and thus cell shrinkage. Indeed, the effect could be transiently reversed when Ca2+ entry was blocked with CdCl2 (Fig. 4D). This suggests that ouabain-induced cell shrinkage contributes to the IOS. In addition, complimentary factors other than changes in total cell volume might participate in the ouabain-induced IOS response (Buchheim et al. 1999; Muller and Somjen 1999). For instance, ouabain-induced excitotoxicity, as described by Zeevalk and Nicklas (1996), might lead to irreversible changes in dendritic morphology, denominated dendritic beading (Andrew et al. 1999; Polischuk et al. 1998). Previously it has been suggested that dendritic beading reflects damage of dendrites, e.g., following a prolonged exposure to hypoxia and/or to high levels of NMDA (Hori and Carpenter 1994; Park et al. 1996). Dendritic beading is expected to result in a decrease in IOS (Andrew et al. 1999; Polischuk et al. 1998). Such action could also account for the irreversibility of the ouabain action.

It can be concluded that an ouabain-induced block of the Na+/K+ pump does not mimic the hypoxic IOS increase. Nevertheless, IOS responses elicited during repetitive hypoxic episodes were blocked in the presence of ouabain. However, ouabain elicited a higher IOS response and a slower recovery after reoxygenation. This can be explained by failure of Na+/K+ pump activity, which reduces the capability of the cells to downregulate their volume. Subsequent hypoxic episodes did not cause IOS responses, which we attributed to severe cell damage (Lees and Leong 1995; Lees et al. 1990).

Further investigations were carried out to ascertain that Na+ influxes through TTX-sensitive Na+ currents contribute to the hypoxic cell swelling. Application of TTX led to a steady decrease in the IOS baseline. This can be attributed to its blockade of neuronal activity, which would lead to a decrease in both neuronal and glial cell volumes. Similar results were reported in the region of the nucleus tractus solitarius in brain stem slices when neuronal activity was blocked (Torres et al. 1997).

Involvement of mitochondrial KATP channels in IOS generation

Another mechanism associated with IOS is swelling of cytoplasmatic organelles, such as mitochondria (Aitken et al. 1999), which has long been known to result of hypoxia and/or ischemia (Aitken et al. 1999; Allen et al. 1989; Lazriev et al. 1980; Vladimirov Iu and Kogan 1981). Swelling of mitochondria is accompanied by a decrease in light scattering (Mar 1981) or light absorbance (Stoner and Sirak 1969). Flow cytometry analysis also showed that swelling of individual mitochondria leads to a decrease in light absorbance (Beavis et al. 1985; Macouillard-Poulletier de et al. 1998), indicating that mitochondria behave as light-scattering objects that affect the IOS in the same way as the whole cell (i.e., swelling leads to an increase in IOS, shrinkage to a decrease).

An indication for the contribution of mitochondrial swelling to IOS generation was observed when KATP channel drugs were applied. KATP channels contribute to IOS in a dual manner. 1) KATP channels regulate the excitability of respiratory neurons (Pierrefiche et al. 1997), and therefore application of KATP channel blockers/activators leads to a rise/fall in the IOS, as the neurons are depolarized/hyperpolarized. In the present experiments, the blocker of plasmalemmal KATP channels, HMR1098, exhibited this behavior. 2) A secondary effect of the drugs seems to target at other structures, e.g., mitochondrial KATP (mitoKATP) channels. Activation of mitoKATP channels with diazoxide was previously observed to induce depolarization of the mitochondrial membrane potential (Grimmsmann and Rustenbeck 1998; Gross and Fryer 1999; Holmuhamedov et al. 1998), which normally induces mitochondrial swelling. KATP antagonists reversed this effect (Garlid et al. 1997). To test for the dual effects of KATP channels, slices were first exposed to a prolonged period of hypoxia prior to drug application to induce maximal activation of plasmalemmal KATP channels. Under such conditions, application of KATP-directed drugs induced IOS responses that could be attributed primarily to changes in mitoKATP channel activity (fall after glibenclamide, rise after diazoxide). The results were remarkably reproducible, indicating that the effects of plasmalemmal and mitochondrial KATP channels indeed differ and can be distinguished in this way. The contribution of mitochondrial IOS signals was further verified through applying the uncoupler of mitochondrial oxidative phosphorylation, CCCP, which depolarizes mitochondria and induces mitochondrial swelling (Minamikawa et al. 1999). Such findings indicate that during prolonged hypoxia, KATP channel blockers and openers act on mitochondrial KATP channels rather than on plasmalemmal KATP channels. A similar finding, i.e., that glibenclamide is much less potent in inhibiting plasmalemmal KATP channels after metabolic poisoning, was previously observed in pancreatic beta -cells (Mukai et al. 1998).

In conclusion, the hypoxic IOS response can be interpreted as a result of several distinct mechanisms as illustrated in Fig. 6. First, there is a component that is related to cell swelling, and, second, there are mechanisms such as dendritic beading or mitochondrial swelling. Cell swelling can be attributed to anoxic depolarization due to hypoxic release and accumulation of neurotransmitters and neuromodulators, while Na+/K+ pump activity is inhibited. The processes acting through Na+ fluxes can be subdivided into two components, one being mediated through voltage-regulated and TTX-sensitive Na+ channels and another that is TTX insensitive.



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Fig. 6. Possible components of the hypoxic IOS response. Boxes depict the qualitative contribution of each mechanism. Note that the IOS component that was not due to cell swelling (denoted "other") was arbitrarily assumed to represent an increase in IOS (as a result of mitochondrial swelling, for instance), whereas it might equally well reflect a decrease (as might occur during dendritic beading).


    ACKNOWLEDGMENTS

We thank N. Hartelt and J. Huhnold for expert technical assistance.


    FOOTNOTES

Address for reprint requests: M. Haller, Physiologisches Institut, Georg-August-Universität Göttingen, Humboldtallee 23, D-37073 Gottingen, Germany (E-mail: mirjam{at}neuro-physiol.med.uni-goettingen.de).

Received 31 July 2000; accepted in final form 14 March 2001.


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0022-3077/01 $5.00 Copyright © 2001 The American Physiological Society




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