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(Stroke. 1996;27:957-964.)
© 1996 American Heart Association, Inc.

Articles

23Na Nuclear Magnetic Resonance Spectral Changes During and After Forebrain Ischemia in Hypoglycemic, Normoglycemic, and Hyperglycemic Rats

Randy L. Tyson, MSc; Garnette R. Sutherland, MD James Peeling, PhD

From the Departments of Chemistry (R.L.T., J.P.), Pharmacology and Therapeutics (J.P.), and Radiology (J.P.), University of Manitoba; the Department of Chemistry, University of Winnipeg (Manitoba) (J.P.); and the Department of Neurosurgery, Foothills Hospital, Calgary, Alberta (G.R.S.), Canada.


*    Abstract
up arrowTop
 *Abstract
down arrowIntroduction
 down arrowMaterials and Methods
 down arrowResults
 down arrowDiscussion
 down arrowReferences
 
Background and Purpose The severity of brain injury in animal models of forebrain ischemia increases with blood glucose level. During ischemia, energy failure is slower and maintenance of ion gradients is prolonged as the level of glycemia increases. It is not clear how the level of glycemia influences recovery of ion homeostasis on reperfusion. It has been shown that changes in the intensity of the multiple-quantum 23Na nuclear magnetic resonance (NMR) signals reflect changes in intracellular Na+ levels. We have used 23Na NMR spectroscopy to evaluate the influence of the level of glycemia on changes in Na+ concentration during and after forebrain ischemia in rats.

Methods Single-quantum (SQ) and double-quantum (DQ) 23Na NMR spectra were measured before and during 10-minute forebrain ischemia and during reperfusion in hypoglycemic, normoglycemic, and hyperglycemic rats.

Results The DQ 23Na NMR signal increased to 210% of preischemia intensity in all rats, but a delay in this increase was observed in normoglycemic and hyperglycemic animals. The rate of the DQ 23Na NMR signal increase was fastest in hypoglycemic (apparent first-order rate constant 0.673±0.046 min-1, P<.002 compared with normoglycemic animals) and slowest in hyperglycemic (0.285±0.024 min-1, P<.03) rats. During reperfusion, the signal intensity recovered rapidly in hypoglycemic (0.385±0.050 min-1) and normoglycemic (0.464±0.047 min-1) rats, whereas in hyperglycemic animals recovery was slow (0.108±0.044 min-1, P<.0001 compared with normoglycemic animals). The SQ 23Na NMR signal intensity increased to 117% of preischemia level in hypoglycemic (P<.05 compared with normoglycemic animals) and to 107% in normoglycemic and hyperglycemic animals during reperfusion.

Conclusions The slower increase in the 23Na DQ NMR signal intensity during forebrain ischemia in rats with higher blood glucose levels suggests that Na+ homeostasis is maintained longer in these animals. On reperfusion, the slower recovery of the DQ 23Na NMR signal intensity in hyperglycemic animals likely indicates a slower recovery of Na+ homeostasis, perhaps contributing to the increased neuronal injury after cerebral ischemia in hyperglycemic animals.


Key Words: cerebral ischemia, transient • glucose • sodium • spectroscopy, nuclear magnetic resonance • rats


*    Introduction
up arrowTop
 up arrowAbstract
 *Introduction
down arrowMaterials and Methods
 down arrowResults
 down arrowDiscussion
 down arrowReferences
 
Approximately 40% of anaerobic metabolism in resting brain is dedicated to the maintenance of ion homeostasis,1 which enables the transmission of neuronal impulses and the maintenance of cell volume, substrate transport, and other processes. The Na+ gradient, maintained by Na+/K+ ATPase, is of particular importance because this form of potential energy is used to maintain Ca2+ homeostasis through the Na+/Ca2+ exchanger, to maintain optimal intracellular pH through the Na+/H+ antiport, and to cotransport glucose and amino acids across the cell membrane. During cerebral ischemia, high-energy phosphates are rapidly depleted, so ion gradients can no longer be maintained and efflux of K+ into the extracellular space and influx of Na+, Ca2+, Cl-, and HCO3- into the intracellular space occurs.

The length of time that ion gradients are maintained during ischemia increases with the level of glycemia,2 3 probably because of the larger amounts of glucose available for anaerobic ATP production. The level of glycemia also affects the extent of brain injury, which is increased by hyperglycemia in models of focal4 5 6 7 8 9 10 and global11 12 13 14 15 ischemia and decreased by moderate hypoglycemia.5 16 17 18 19 Increased ischemic damage is therefore not due to events triggered by faster failure of ion gradients during short-duration ischemia. However, the extent of injury may be related to the recovery of ion homeostasis in the reperfusion period. No differences are observed in the reperfusion recovery of extracellular K+, Ca2+, and H+ concentrations after ischemia in normoglycemic and hyperglycemic animals, although electroencephalographic recovery is slower after hyperglycemic ischemia.3 Recovery of intracellular ion concentrations during reperfusion in relation to blood glucose remains unstudied.

NMR relaxation of 23Na nuclei (nuclear spin 3/2) in biological environments occurs in a biexponential fashion through the interaction of these nuclei with charged polyelectrolytes (proteins); the application of a DQ filter allows the observation of Na+ in environments that contain a high concentration of polyelectrolytes.20 Because such high concentrations of proteins occur in the intracellular compartment, the observed 23Na DQ NMR signal of tissue arises mainly from intracellular sources, and a growing body of literature suggests that changes in the intensity of multiple-quantum 23Na NMR signals parallel changes in the intracellular Na+ concentration.21 22 23 24 25 26 27 In the present study, the influence of the level of glycemia on changes in the SQ and DQ 23Na NMR spectra during and after transient forebrain ischemia were evaluated.


*    Materials and Methods
up arrowTop
 up arrowAbstract
 up arrowIntroduction
 *Materials and Methods
down arrowResults
 down arrowDiscussion
 down arrowReferences
 
All animals were cared for in accordance with the guidelines of the Canadian Council on Animal Care. Male Sprague-Dawley rats weighing 250 to 320 g were fasted for 24 hours before use in all experiments. Each rat was treated with atropine (0.5 mg/kg), anesthetized with sodium pentobarbital (60 mg/kg), intubated, and mechanically ventilated throughout the experiment. Brain temperature was monitored throughout the experiment with a tympanic membrane thermocouple probe28 (Cole-Parmer) and maintained at 38.2±0.2°C. The tail artery was exposed and catheterized to allow samples of blood to be taken for blood gas, osmolarity, and blood glucose measurement and for blood pressure recording throughout the experiment. To allow administration of glucose and insulin solutions, the femoral vein was catheterized. The carotid arteries were exposed through a midline neck incision, and snares were placed around them in preparation for occlusion during ischemia. The scalp and temporalis muscles were removed (bleeding being prevented by electrocauterization), and a surface coil for the reception of NMR signals was positioned on the exposed cranium.

Each rat was placed into an animal holder, the femoral catheter was connected to an infusion pump (Sage syringed pump, model 355, Cole-Parmer), and blood glucose was altered to the desired level (see below). The animal holder was placed into the magnet, and preischemia 23Na NMR spectra were obtained as described below.

After the acquisition of the preischemia spectra, blood pressure was lowered to a mean of 45 mm Hg through aspiration of blood from the tail catheter into a heparinized syringe, at which point the carotid arteries were occluded. Acquisition of 31P or 23Na NMR spectra was begun at the time of occlusion. After 10 minutes of ischemia, cerebral perfusion was restored, and the aspirated blood was reinfused. In normoglycemic animals, forebrain ischemia of this duration results (after a 7-day interval) in extensive neuronal necrosis in the CA1 sector of the hippocampus and the dorsal-lateral striatum, and in a mild to moderate degree of necrosis among pyramidal neurons in midneocortical layers.29

Blood glucose was monitored with a blood glucose meter (One Touch, Lifescan Canada Ltd), with measurements based on the glucose oxidase method for whole blood.30 Before ischemia, blood glucose was adjusted to one of three groups: hypo (blood glucose {approx}2 mmol/L, n=7); normo (blood glucose {approx}6 mmol/L, n=7); or hyper (blood glucose {approx}20 mmol/L, n=7). Blood glucose levels were altered by administration of boluses and infusions (titrated against glucose measurements taken every 10 minutes) as described below. The hypo and normo groups were made iso-osmolar with the hyper group by including 25% wt/vol mannitol in the solutions. A fourth group (blood glucose {approx}7 mmol/L, n=7), used as a control to enable the observation of possible effects due to changes in osmolarity, was kept normoglycemic without the addition of mannitol. Animals were stabilized at their preischemic blood glucose levels for 20 minutes before ischemia.

In the hypo group, hypoglycemia was induced with a bolus of 0.20 mL·100 g-1 of 7 IU/mL insulin and 25% wt/vol mannitol/saline (delivered over a 30-second interval to minimize physiological effects, eg, to the blood-brain barrier) followed by an infusion of 0.5% wt/vol glucose and 25% wt/vol mannitol/saline at a rate of 0.98±0.15 mL·100 g-1·h-1. Rats in the normo group were given a 0.20-mL·100 g-1 bolus of 5% wt/vol glucose and 25% wt/vol mannitol/saline, followed by infusion of the same solution (mean rate, 1.10±0.07 mL·100 g-1·h-1, titrated to a blood glucose concentration of 6 mmol/L). Hyperglycemia in the hyper group was produced by giving each animal a 0.20-mL·100 g-1 bolus of 65% wt/vol glucose/saline followed by infusion of the same solution (mean rate, 1.00±0.28 mL·100 g-1·h-1, titrated to a blood glucose concentration of 20 mmol/L). The control group was given a 0.20-mL·100 g-1 bolus and infusion (mean rate, 1.16±0.18 mL·100 g-1·h-1, titrated to a blood glucose concentration of 6 mmol/L) of 5% wt/vol glucose/saline with no mannitol for adjustment of osmolarity.

To determine the dependence of the intensity of the 23Na DQ NMR signal on pH, samples of homogenized brain tissue (n=5) were prepared and titrated with small amounts of 1 mol/L NaOH/deuterated saline or 1 mol/L HCl/deuterated saline over the range of pH 7.3 to pH 5.5 at 38°C. The titration was started at pH 5.5 for three samples and at pH 7.3 for two samples so that any systematic changes in samples over time could be detected. At each pH sampled, 31P and SQ and DQ 23Na NMR spectra were obtained.

All in vitro 79.39-MHz 23Na NMR experiments were performed at 38.0°C with a Bruker AM300 NMR spectrometer (Bruker Analytische Messtechnik GMBH) with a commercial multinuclear broadband probe. Shimming was performed using the 1H NMR signal from water. Each in vitro SQ 23Na NMR spectrum consisted of the sum of 64 transients (1024 data points) with spectral width of 5000 Hz, acquisition time of 0.050 s, and relaxation delay of 0.085 s. The spectra were collected using CYCLOPS phase cycling. Both in vitro and in vivo (see below) DQ-filtered 23Na NMR spectra were obtained using the sequence31 90°{phi}–{tau}/2–180°{phi}±90°–{tau}/2–90°{phi}+90°–{delta}–90°–acquire, where {phi} is the axis along which the excitation field B1 is applied in the rotating frame of reference, {tau} is the echo delay time, and {delta} is the DQ evolution time. The DQ 23Na NMR signal was separated from SQ and triple-quantum signals also generated by this pulse sequence, using a 32-step phase cycling procedure.32 No significant changes in the signal intensity of the 23Na SQ resonance occurred in any sample throughout the experiment, so changes in intensity of the 23Na DQ resonance directly reflected the effect of changes in pH. Spectral acquisition parameters for the in vitro DQ 23Na NMR spectra were similar to those used for the in vitro SQ 23Na NMR spectra, with {tau} selected to be 8 milliseconds (the optimized value of {tau} used in the in vivo studies as described below).

Sample pH was determined from the difference in chemical shift of inorganic phosphate and phenylphosphonate (added to the NMR samples to serve as an internal reference) from 121.3-MHz 31P NMR spectra, and this value was compared with the corresponding pH from a previously prepared calibration curve (data not shown). Spectra were obtained for 8 to 15 pH values for each sample.

A linear fit to the plot of 23Na DQ peak height against pH yielded the change in the intensity of the 23Na DQ resonance per pH unit, defined as fpH. The value of fpH was used to correct the in vivo DQ 23Na NMR signal intensity for the effects of pH determined as outlined below during and after ischemia.

Using published in vivo 31P NMR data,33 we fitted plots of tissue pH against time during ischemia for each blood glucose group to the data with an equation of the form34

(1)

where the superscript i represents ischemia, the subscripts 0 and {infty} indicate the values of pH at the start of ischemia and at infinity, respectively, and kpHi is the apparent first-order rate constant for the exponential decrease in pH during ischemia. The data obtained after ischemia were fit to the equation

(2)

where the superscript p represents after ischemia, pH0 and pH{infty} are the values of pH at the beginning of reperfusion and at an infinite time of reperfusion, and kpHp is the apparent first-order rate constant for the recovery of pH during reperfusion. Thus, at any time point the change in pH, {Delta}pH, can be calculated and used to correct the DQ signal intensity for the effects of pH changes.

Interleaved in vivo SQ and DQ 23Na NMR spectra were obtained at 79.455 MHz with a Bruker Biospec 7/21 spectrometer (Bruker Analytische Messtechnik GMBH). For transmission, a 5-cm diameter saddle coil tuned to both the 1H (300.13 MHz) and 23Na resonance frequencies was used, while reception was accomplished through the use of a 1.8x2.3-cm elliptical surface coil tuned to the 23Na frequency and orthogonalized with respect to the saddle coil. We have shown previously that a correctly positioned surface coil of this size effectively samples the forebrain with minimal detection of signals from adjacent muscle.29

The SQ 23Na NMR spectra were obtained using a one-pulse sequence, each consisting of the sum of 24 transients (512 data points) collected with CYCLOPS phase cycling, with preacquisition delay of 100 µs, relaxation delay of 0.085 s, spectral width of 2500 Hz, and acquisition time of 0.1024 s.

Each DQ 23Na NMR spectrum (512 data points) consisted of 128 scans and acquisition parameters identical to those for the SQ 23Na NMR spectra, and with the evolution time {delta} set to 25 µs and the echo delay time {tau} set to 8 ms. The DQ 23Na NMR signal was found to be at a maximum for this value of {tau}. The 90° pulse was 280 µs. Acquisition of an SQ and DQ 23Na NMR spectral pair required 30 s; for each rat, 10 pairs of interleaved SQ and DQ 23Na NMR spectra were obtained before ischemia, 20 pairs during ischemia, and 120 pairs after ischemia.

A 5-Hz line-broadening function was applied to the SQ 23Na NMR free-induction decays after zero-filling to 2048 data points. The free-induction decays were Fourier transformed, and the resulting spectra were phased. Baseline distortion in each spectrum was removed using a deconvolution baseline correction procedure. Peak integrals were measured relative to the mean preischemia level.

Each DQ 23Na NMR free-induction decay was multiplied with a 25-Hz line-broadening function to optimize signal-to-noise ratio after zero-filling to 2048 data points. After Fourier transformation of each free-induction decay, the resulting spectrum was phased, and the peak height relative to the mean preischemia peak height was used in the analysis. Signal-to-noise ratio was insufficient to apply the methods of line-shape simulation to the DQ 23Na spectra. Any changes in the relaxation parameters of the DQ resonance would be negligible to the line-broadening function, and measurement of peak height is affected less than integrated intensity by errors due to phase and baseline correction.35 The raw DQ 23Na NMR peak heights were then multiplied by 1/(1-fpH·{Delta}pH), where fpH is the change in the 23Na DQ NMR peak height per pH unit (above) and {Delta}pH is the difference in pH relative to the preischemia pH at each time point during ischemia and reperfusion, to obtain the pH-corrected 23Na DQ NMR peak heights.

Equation 3Down was used to fit the pH-corrected 23Na DQ peak height data obtained during ischemia:

(3)

where ti is the time from the start of ischemia, Ii and Ii{infty} are the DQ 23Na NMR signal peak heights at ti=0 and ti={infty}, respectively, and kDQi is the apparent first-order rate constant for the increase in the peak heights during ischemia. Because a delay of approximately 30 s was observed before an increase in the DQ 23Na NMR peak height occurred in the normo and hyper groups, ti=0 was set at 30 s of ischemia for these groups.

The DQ peak height fell to below the preischemia value early in reperfusion in the hypo and normo groups, with subsequent recovery. Therefore, to fit the data obtained during reperfusion, a third term was added to give an equation of the form

(4)

where tp is the time from the start of reperfusion, Ip is the DQ 23Na NMR peak height relative to the preischemia peak height at time tp, I0p and I{infty}p are the peak heights at tp=0 and tp={infty}, respectively, Irecp represents the extent of the undershoot in the DQ 23Na NMR peak height early in reperfusion, kDQp is the apparent rate constant for the initial fall in the DQ peak height, and kDQrec is the apparent rate constant for the recovery from the undershoot.

Physiological variables (levels of blood gases, osmolarity, hematocrit, blood glucose, and blood pressure) were compared among groups using ANOVA, with post hoc comparisons made using Scheffé's method of intergroup comparison (level of significance, P<.02).

The temporal curves of SQ and DQ 23Na NMR intensity data from the control, hypo, normo, and hyper groups were compared using repeated measures least-squares analysis (level of significance, P=.05). For those curves showing significant interaction and P>.02, data points were compared (with Bonferroni correction) at each time point. Parameter estimates from Equations 3 and 4 were obtained using a Levenberg-Marquardt nonlinear least-squares minimization algorithm. The level of statistical significance corresponded to the probability value giving largest confidence limits without overlap of the intervals for parameters being compared.


*    Results
up arrowTop
 up arrowAbstract
 up arrowIntroduction
 up arrowMaterials and Methods
 *Results
down arrowDiscussion
 down arrowReferences
 
Table 1Down gives the physiological data measured during this study. The difference in preischemia plasma osmolarity between the control and hyper groups was on the verge of significance (P=.089) and was significant at 20 (P=.0061) and 60 minutes (P=.0056) of reperfusion. No significant differences in preischemic or postischemic hematocrit levels, blood pH, PCO2, PO2, or HCO3- were observed.


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  		Table 1. Physiological Variables for Each Blood Glucose Group Before and After Transient Forebrain Ischemia

Fig 1Down shows the SQ 23Na NMR integrated signal intensity for the hypo, normo, and hyper groups during and after transient forebrain ischemia. No significant differences were observed between the control and normo groups either during ischemia (P=.53) or reperfusion (P=.73), so the control group is not shown. During ischemia, the apparent decrease in the SQ 23Na NMR intensity was not statistically significant. After reperfusion, the signal intensity of the SQ 23Na NMR resonance in the normo and hyper groups increased to 107% of preischemia signal intensity at the end of the 1-hour reperfusion period. The hypo group showed a significantly greater increase in signal intensity than the normo and hyper groups (117% relative to preischemia; P=.0060 compared with the hyper group, P<.05 from 20 to 60 minutes of reperfusion compared with the normo group).



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  		Figure 1. Signal intensity (mean±SD) of the SQ 23Na NMR resonance during and after ischemia in hypoglycemic, normoglycemic, and hyperglycemic rats. Shaded area indicates the period of ischemia.

The in vitro experiments using homogenized brain showed that the peak height of the 23Na DQ NMR resonance decreased 23±3% per unit decrease in pH over the range studied. Thus, the intensity of the observed 23Na DQ NMR signal during ischemia was significantly altered, and to different extents in different blood glucose groups, because pH fell below 5.8 in the hyper and to only 6.7 in the hypo group in this model after 10 minutes of ischemia.33

The time-series raw (pH-uncorrected) DQ 23Na NMR data for the hypo, normo, and hyper groups are given in Fig 2Down; the pH-corrected results are shown in Fig 3Down. The data curves displayed marked differences among blood glucose groups (P<.001). Data for the control group are not included because 23Na DQ NMR spectra from the normo and control groups showed no differences, either during or after ischemia. Any physiological perturbations that might be introduced by the use of mannitol to maintain uniform osmolarity among the groups were apparently insufficient over the course of the 90-minute experiment to affect the NMR spectra.



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  		Figure 2. Changes in the observed DQ 23Na NMR peak heights (mean±SD) during and after transient cerebral ischemia in hypoglycemic, normoglycemic, and hyperglycemic rats. Shaded area indicates the period of ischemia.



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  		Figure 3. pH-Corrected DQ 23Na NMR peak heights (mean±SD) from Fig 2Up. Shaded area indicates the period of ischemia, and curves are best fits to the data using Equations 3 and 4 with parameter values listed in Table 2Up.

The apparent first-order rate constant kDQi was significantly different among groups and graded with respect to the level of glycemia (Table 2Down), being greatest in the hypo group (P<.002 compared with normoglycemia and P<.0005 compared with hyperglycemia) and least in the hyper group (P<.03 compared with normoglycemia). In the hypo group, the DQ signal intensity rose within the first 30 s of ischemia, reaching 200% of the preischemic level by 120 s and slowly increasing to 225% by the end of 10 minutes of ischemia. In both the normo and hyper groups during ischemia, a delay of about 30 s was observed before the DQ signal intensity increased, reaching 210% of preischemic level at 10 minutes of ischemia. The parameter I{infty}i shows that the DQ 23Na NMR signal intensity for all groups would reach about 220% of preischemia at long ischemia times.


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  		Table 2. Results of Curve Fit to pH-Corrected DQ 23Na NMR Peak Height Data in Fig 3Up Using Equations 3 and 4

After ischemia, the DQ 23Na NMR signal intensity in the hypo and normo groups decreased to 85% of the preischemic level after about 7 minutes and recovered to about 95% of preischemic level by the end of the experiment. The total recovery, given by I{infty}p+Irecp, for the hypo and normo groups was not significantly different from 1.0, indicating that complete recovery would occur at a time longer than that studied. Recovery of the DQ 23Na NMR signal was rapid during early reperfusion in the hypo and normo groups, the apparent first-order rate constant for recovery being greater in the normo group (kDQp=0.385 min-1 for the hypo group, kDQp=0.464 min-1 for the normo group; P=.15). In marked contrast to the lower blood glucose groups, recovery of the DQ signal intensity in the hyper group was much slower (kDQp=0.108 min-1; P<.001 versus hypo and normo groups). The peak height remained slightly elevated at the end of 1 hour of reperfusion, and the results of the curve fit in Table 2Up indicate that recovery would not occur at longer times. Neither the values of Irecp nor kDQrec were significantly different among the three groups.


*    Discussion
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 up arrowAbstract
 up arrowIntroduction
 up arrowMaterials and Methods
 up arrowResults
 *Discussion
down arrowReferences
 
The increase in Na+t concentration after 10 minutes of forebrain ischemia observed in the present study agrees well with increases found using a four-vessel occlusion model of global ischemia of 15-minute duration36 and using a temporary middle cerebral arterial occlusion model of 1-hour duration.37 The increase in Na+t after ischemia may be due to increased leakage across the blood-brain barrier due to stimulation of Na+/K+ ATPase in the endothelial cells of microvessels,38 thereby transferring Na+ (and water) from blood to the extracellular compartment. It has been suggested that free fatty acids and free radicals generated during ischemia may contribute to such a stimulation of Na+/K+ ATPase.39 Glucose, on the other hand, inhibits endothelial Na+/K+ ATPase,40 41 so postischemic stimulation may be more pronounced in hypoglycemic rats, leading to a greater increase in Na+t in these animals. This may explain the greater increase in the SQ 23Na NMR signal intensity after ischemia observed in hypoglycemic rats in this study.

The 23Na nucleus has a nuclear spin quantum number of 3/2. Under conditions where the motional correlation time is of the order of the inverse of the Larmor frequency, biexponential relaxation can result.20 Restricted motion satisfying this condition can be caused by interaction of Na+ with charged macromolecules (polyelectrolytes) such as proteins. Biexponential relaxation of quadrupolar nuclei then allows the detection of multiple-quantum coherence signals in tissue.42 There is evidence that in perfused organs changes in the multiple-quantum 23Na NMR signal correspond to changes in intracellular sodium concentration. Thus, the addition of acetylcholine in the presence of ouabain to perfused rat salivary gland causes a large increase in intracellular Na+, with a concurrent increase in the DQ 23Na NMR signal.26 In perfused kidney, both anoxia and ischemia produce large increases in the DQ 23Na NMR signal,22 and addition of ouabain to the perfusate produces large increases in the triple-quantum 23Na NMR signal.23 Ischemia in the perfused heart produces large increases in intracellular Na+ as measured by SQ 23Na NMR spectroscopy in the presence of a shift reagent, which separates the NMR signals from intracellular and extracellular Na+, with a concurrent increase in the DQ 23Na NMR signal.25 Indeed, at short DQ preparation times {tau} (<=4 ms), in perfused liver the DQ 23Na NMR signal is due almost exclusively to intracellular sodium.27 There is also evidence that changes in DQ 23Na NMR signal reflect changes in intracellular sodium concentration in brain. Following the addition of veratridine (a Na+-channel agonist) to brain slices, a large increase in the DQ 23Na NMR signal intensity occurs due to increased intracellular Na+ following the opening of Na+ channels.24 In the presence of tetrodotoxin, veratridine has no effect on the DQ 23Na NMR signal.24 At death, a large increase in DQ 23Na NMR signal and a decrease in the DQ 39K NMR signal are observed,21 consistent with the movement of Na+ into and K+ out of the intracellular space.

Changes in the DQ 23Na NMR signal due to ischemia and reperfusion then reflect movement of Na+ between the extracellular and the intracellular compartments. The values in Table 2Up for the apparent first-order rate constants kDQi and kDQp demonstrate that there is a clear dependence in the rates of movement of Na+ on the level of glycemia. During ischemia, kDQi is smallest in hyperglycemic and largest in hypoglycemic animals, and in addition the onset of the increase in the DQ 23Na NMR signal is delayed in the higher blood glucose groups. These observations indicate that there is a slower and delayed increase in the intracellular Na+ concentration during ischemia in the setting of elevated blood glucose levels. The concentrations of the high-energy phosphates PCr and ATP similarly decrease at rates that decrease with increasing blood glucose level.33 These observations are consistent with prolonged maintenance of Na+ homeostasis due to ongoing Na+/K+-ATPase activity, supported by ATP produced through anaerobic glycolysis of elevated brain glucose stores, during ischemia in rats with higher blood glucose levels. A similar glucose-dependent delay in the increase of extracellular K+ during ischemia has been observed using ion-selective microelectrodes.2 3

With reestablished blood flow, the DQ 23Na NMR signal rapidly decreases in intensity in the hypo and normo groups (kDQp=0.385 and 0.464 min-1, respectively), consistent with rapid movement of Na+ out of the intracellular space. However, under hyperglycemic conditions, recovery is significantly slower (kDQp=0.108 min-1). This parallels the much slower recovery of ATP during reperfusion in hyperglycemic animals,33 43 so the slower recovery of Na+ homeostasis may be due in part to prolonged suppression of Na+/K+-ATPase activity that follows hyperglycemic ischemia. However, in addition there is a large increase in the intracellular H+ concentration during ischemia in hyperglycemic animals10 33 43 44 and slower recovery of normal pH after reperfusion.33 The elevated intracellular H+ levels may contribute to the slow recovery of intracellular Na+ by stimulation of the Na+/H+ antiport,45 countering the removal of Na+ from the intracellular space by Na+/K+-ATPase.

The I{infty}i values for the three blood glucose groups are similar, consistent with intracellular Na+ reaching a similar concentration during ischemia in all groups. The maximum change in the DQ signal attained during ischemia is similar to that obtained in hypoxic brain slices,24 and the 220% increase (I{infty}i in Table 2Up) agrees well with a 205% increase in the intracellular concentration of Na+ measured using microelectrodes in rat cortex during global ischemia.46

After reperfusion, the DQ 23Na NMR signals in the hypo and normo groups fall below the preischemic level before complete recovery, as given by the value of I{infty}p in Table 2Up. The significance of the Na+ undershoot in the hypo and normo groups early in reperfusion is not clear, but similar undershoots involving extracellular K+ concentrations have been observed after ischemia47 48 and neuronal excitation49 and have been attributed to enhanced postischemic activity of Na+/K+ ATPase50 51 in these cases. After reperfusion in hyperglycemic animals, however, the DQ Na23 NMR signal returns much more slowly to the preischemic level with no undershoot, consistent with the argument above that delayed recovery of ATP levels may impede the resumption of Na+/K+ ATPase activity in these rats. This slower recovery of intracellular Na+ following ischemia may be an important factor contributing to the accentuated ischemia brain injury in the presence of elevated blood glucose levels. For instance, Na+ carries water as it is transferred across cell membranes, so the slower removal of intracellular Na+ may contribute to the observed increase in postischemic cytosolic edema in hyperglycemic animals.52 53 54 Furthermore, slower recovery of Na+ homeostasis may impede the function of the Na+/Ca2+ exchanger,55 a major transporter of Ca2+ from the intracellular space, and contribute to the observed slower recovery of Ca2+ after ischemia in hyperglycemic animals.56

Changes in the SQ 23Na NMR signal intensity have been used to follow Na+ movement between the intracellular and extracellular spaces during generalized seizure in cats57 and global cerebral ischemia in dogs.58 The present study shows that, whereas small changes in SQ 23Na NMR signal intensity occur during ischemia, the DQ 23Na NMR signal intensity is a much more sensitive monitor of changes in intracellular Na+ concentration during and after ischemia.


*    Selected Abbreviations and Acronyms
 
DQ = double-quantum
hyper group = hyperglycemic group
hypo group = hypoglycemic group
Na+t = brain tissue sodium
NMR = nuclear magnetic resonance
normo group = normoglycemic group
SQ = single-quantum


*    Acknowledgments
 
This study was supported by the National Research Council of Canada and the Heart and Stroke Foundation of Canada. The authors thank Maureen Donnelly, Fang Wei Yang, and Yan Fan for their assistance in animal preparation.


*    Footnotes
 
Reprint requests to Dr James Peeling, Department of Pharmacology and Therapeutics, University of Manitoba, 770 Bannatyne Ave, Winnipeg, Manitoba, Canada R3E 0W3. E-mail jim@bionmr.mrrl.umanitoba.ca.

Received September 28, 1995; revision received January 30, 1996; accepted January 30, 1996.


*    References
up arrowTop
 up arrowAbstract
 up arrowIntroduction
 up arrowMaterials and Methods
 up arrowResults
 up arrowDiscussion
 *References
 

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