Glycemia-dependent Nuclear Factor κB Activation Contributes to Mechanical Allodynia in Rats with Chronic Postischemia Pain

ISCHEMIA–REPERFUSION (I/R) injury involves a prolonged period of ischemia followed by reperfusion, leading to both deleterious intracellular injuries and inflammatory reactions.¹  Postischemia pain is a well-recognized consequence of I/R injury,²  and chronic postischemia pain (CPIP) is an animal model of long-term postischemia pain. Under general anesthesia, CPIP rats undergo placement of a tight-fitting tourniquet around their hind paw for a 3-h period followed by reperfusion; chronic ipsilateral and more sporadic contralateral, mechanical, and cold allodynia were demonstrated to follow this I/R injury for at least 4 weeks.³  We have previously demonstrated that glycemic modulation during I/R injury can significantly impact allodynia in CPIP rats. Indeed, hyperglycemia (dextrose administration) during I/R injury led to significantly increased mechanical and cold allodynia for at least 3 weeks after the injury, whereas, relative hypoglycemia (fasting and insulin administration) resulted in significantly decreased mechanical and cold allodynia.⁴ 

Nuclear factor κB (NFκB) is an inducible redox-sensitive transcription factor present in a latent form in the cytoplasm of almost all mammalian cells.^5–7  It regulates many aspects of cellular stress and initiates the transcription of several proinflammatory mediators, such as cytokines, adhesion molecules, and inflammatory enzymes.⁸  NFκB activation can be modulated by different glycemic conditions. Indeed, NFκB is activated in tissues exposed to chronic high glucose.⁹  Chronic high glucose leads to increased reactive oxygen species and tumor necrosis factor-α, which mediate the degradation of inhibitor κB-α, a regulator of NFκB, allowing NFκB nuclear translocation. Increases in activated NFκB and tumor necrosis factor-α levels were also demonstrated in healthy subjects after acute ingestion of glucose or a mixed meal.^10,11 

In contrast, insulin has inhibitory effects on NFκB activation. Insulin was shown to inhibit NFκB in human aortic endothelial cells in vitro^12,13  and in monocytes in vivo.¹⁴  Similarly, infusing insulin to obese subjects increases inhibitor κB-α levels, and decreases NFκB.¹⁴  Similar results were obtained in a rat model of type 2 diabetes mellitus, where insulin increases inhibitor κB-α and decreases NFκB p65 in the liver and skeletal muscle of the animals.¹⁵ 

NFκB exacerbates cellular injury and inflammation in animal models of cerebral,¹⁶  spinal cord,¹⁷  and skeletal muscle¹⁸  I/R injury. NFκB may enhance postischemia pain after I/R injury, and therefore, might be an interesting target for intervention. In the current study, we hypothesize that the increased mechanical allodynia observed in CPIP rats with higher glycemic levels is associated with increased NFκB activation, and that conversely, the reduced mechanical allodynia observed in relatively hypoglycemic CPIP rats is associated with reduced NFκB activation. We also hypothesize that NFκB inhibition will significantly reduce postischemia pain, and that groups of CPIP rats with higher glycemic levels will exhibit a rightward shift of the antiallodynic dose–response curve after NFκB inhibition compared with the relatively hypoglycemic group of CPIP rats, reflecting different intrinsic levels of activated NFκB.

The current study was conducted in male Long Evans rats (300–500 g; Charles River, Senneville, Quebec, Canada). After their arrival, rats were allowed a minimum of 3 days to acclimatize before the start of the experiment. Rats were housed in groups of two to four, had unlimited access to food and water, and were exposed to a 12-h light:dark cycle (lights on 7:00 am). Experiments were approved by the McGill Animal Care and Ethics committees (McGill University, Montreal, Quebec, Canada), and were performed in accordance with the ethical guidelines of the Canadian Council on Animal Care (Ottawa, Ontario, Canada) and the International Association for the Study of Pain (Washington, DC).¹⁹ 

I/R injury of the left hind paw was induced, as described.³  Rats were deeply anesthetized with an intraperitoneal bolus (55 mg/kg) of sodium pentobarbital, followed by a 2-h continuous infusion (27.5 mg·kg⁻¹·h⁻¹, intraperitoneal). After anesthesia induction and verification of its depth by confirming the absence of pedal reflexes, a Nitrile 70 Durometer O-ring (O-rings, Seattle, WA) with a 5.5 mm internal diameter was placed around the rat’s left hind paw, proximal to the ankle and left in place for 3 h, after which it was cut to allow reperfusion. The termination of the pentobarbital infusion was timed so that rats would recover within 60 min after reperfusion. Sham surgery consisted of pentobarbital anesthesia alone.

Rats undergoing I/R or sham injury were divided into four treatment groups, in order to evaluate different glycemic conditions. The first group of rats had normal access to food and water (fed; seven CPIP/six sham-treated). The second group of rats had normal access to food and water, but also received subcutaneous injections of insulin diluted in normal saline (fed/insulin; six CPIP/seven sham-treated), according to a sliding scale designed for this experiment (baseline glycemia: ≤6: 0.5 U/kg; >6 ≤7: 1 U/kg; >7 ≤8: 1.5 U/kg; >8 ≤9: 2 U/kg; >9 ≤10: 2.5 U/kg; because none of the rats had a baseline glycemia >10, it was not necessary to extend the insulin sliding scale further). The total dose of insulin was administered in two half-doses, the first one immediately before ischemia, and the second one 60 min after ischemia induction. The maximal volume of insulin solution administered was 50 µl. A third group of rats had normal access to food and water, but also received insulin according to the above-described sliding scale as well as dextrose (fed/insulin/dextrose; six CPIP/eight sham-treated). Dextrose was dissolved in deionized water (final concentration: 40%) and administered as 1 ml intraperitoneal injections at baseline, 60, and 120 min after the induction of ischemia. A fourth group of rats (fed/dextrose; nine CPIP/eight sham-treated) consisted of normally fed animals given dextrose according to the above schedule in order to produce a relative hyperglycemia.

Glycemic levels were measured with a glucometer (FreeStyle mini; Abbott Diabetes Care, Mississauga, Ontario, Canada). Blood was withdrawn from the extremity of the rat tail, from a submillimetric incision, by an observer blinded to treatment. We did not measure local glucose levels because we determined in preliminary trials that sampling blood from the hind limb produced local injury that produced significant allodynia on its own. Volume of blood withdrawn varied between 10.0 and 40.0 μl. Blood samples were withdrawn at baseline, and then 30, 90, and 150 min after the placement of the tourniquet (ischemia phase), as well as 20, 40, and 60 min after the removal of the tourniquet (reperfusion phase).

Insulin R (Eli Lilly Canada Inc., Toronto, Ontario, Canada) was diluted in sterile normal saline to obtain a final concentration of 10 U/ml. SN50 (Enzo Life Sciences International Inc., Plymouth Meeting, PA) was diluted in normal saline to a concentration of 50 ng/μl. Solutions were made freshly before the experiment. The inactive control peptide of SN50 (Enzo Life Sciences International Inc., Plymouth Meeting, PA) stored at −20°C was similarly diluted in normal saline to a concentration of 50 ng/μl, and freshly diluted to a final concentration of 30 ng/μl before the experiment.

Four CPIP and four sham groups of rats–fed (seven CPIP and six sham), fed/insulin (six CPIP and seven sham), fed/insulin/dextrose (six CPIP and eight sham), and fed/dextrose (nine CPIP and eight sham) were tested for mechanical allodynia on day 2 post-I/R procedure by an observer blinded to treatment. Six animals randomly selected from each group (seven from CPIP fed rats) were killed by decapitation after a lethal dose of sodium pentobarbital. Muscle samples (12–30 mg wet weight each) of the superficial plantar layer (flexor hallucis brevis, flexor digiti minimi, and flexor digitorum brevis) and spinal cord samples (10–20 mg wet weight each) at the lumbar (L3–L6) segments, ipsilateral to the tourniquet placement, were immediately extracted. Spinal cord samples were horizontally sectioned in half, to isolate the dorsal part containing the dorsal horns, and then vertically sectioned in half at the midline, to separate the ipsilateral from the contralateral side. Samples were flash frozen in isopentane (−45°C), put on dry ice, and kept at −80°C until processing.

Nuclear extracts were prepared according to a protocol from a commercially supplied kit (Active Motif, Carlsbad, CA). Briefly, tissue samples from the ipsilateral paw muscles (left paw in sham-treated animals) and the ipsilateral dorsal horn (left dorsal horn in sham-treated animals) were diced into very small pieces using a clean razor blade. Pieces were collected in a prechilled container. Tissues were placed on ice and were disrupted and mechanically homogenized in a lysis buffer. Samples were then centrifuged at 850g in a microcentrifuge precooled to 4°C for 10 min. The pellets were then resuspended in a hypotonic buffer, after which they were incubated for 15 min on ice. Detergent was added and the suspension was centrifuged for 30 s at 14,000g in a microcentrifuge precooled to 4°C. The supernatant was stored at −80°C. Nuclear-enriched pellets were resuspended in a lysis buffer and were vortexed for 10 s at maximal intensity. They were then incubated for 60 min on ice, followed by 30 s vortexing at maximal intensity, and then concentrated by centrifugal filtration at 14,000g for 20 min, using cellulose filters with a 30 kDa cutoff (Microcon YM-30; Millipore Corp., Billerica, MA). The resulting nuclear fractions were aliquoted and stored at −80°C until further analysis. Total protein content of nuclear extracts was determined according to the Bradford method.²⁰ 

In order to measure NFκB levels, we used a commercially supplied p65 NFκB transcription factor binding assay (Active Motif), according to the protocol described by the manufacturer. Duplicates of nuclear samples containing 25 μg of total protein (volume 20–30 μl) were deposited in buffer in wells (final volume 150 μl) containing the NFκB consensus site oligonucleotide sequence (5′-GGGACTTTCC-3′) and incubated overnight at 4°C. The N-terminus of NFκB is significantly conserved; consequently, the NFκB consensus motif of the assay binds both human and rat p65. After three washes with 200 μl of wash buffer, 100 μl of diluted rabbit NFκB antibody (1:1,000) was added to each well and incubated for 60 min at room temperature. After three washes with 200 μl of wash buffer, 100 μl of horse radish peroxidase-conjugated antibody (1:1,000) was added to the wells, after which the samples were incubated for 60 min at room temperature. Colorimetric detection was then used to semiquantitatively assess nuclear (translocated) NFκB p65 levels, measured as net absorbance (total absorbance minus background and negative control levels) on a spectrophotometer at 450 nm (Molecular Devices, Sunnyvale, CA).

The NFκB inhibitor SN50 was used to investigate the involvement of NFκB in mechanical allodynia in relatively hyperglycemic CPIP rats versus normo- and relatively hypoglycemic CPIP rats. SN50 is a cell-permeable synthetic peptide, which inhibits the translocation of the active NFκB complex into the nucleus, in cell cultures²¹  and in vivo.²² 

The effects of SN50 on NFκB inhibition were studied in CPIP rats (N = 7 per group) in each of the four glycemic conditions described above (i.e., fed, fed/insulin, fed/insulin/dextrose, and fed/dextrose). Five different drug treatments were tested for each group. Each group of rats received four doses of SN50 intrathecally, every other day (four different SN50 doses between 6.66 and 1,000 ng) and 600 ng of inactive control peptide on the last day of treatment. The first SN50 treatment was administered on day 2 post-IR injury, after testing for mechanical allodynia (baseline score), and the last treatment was given on day 10. Intrathecal injections (all 20 μl) were performed by lumbar puncture at L5 and L6 level. SN50 injections were performed under isoflurane anesthesia (2.5–3.5% in O₂). Rats were then returned in their cages for a 20-min recovery period, after which they were placed in clear acrylic cages (25 × 12.5 cm² with a height of 16 cm), fitted with a stainless steel wire mesh floor for a 10–20 min acclimatization period before allodynia testing. Rats were tested for mechanical allodynia immediately before, and then 30, 75, 120, 165, and 210 min after SN50 administration, by an observer blinded to treatment.

Ipsilateral mechanical allodynia was assessed by measuring paw withdrawal thresholds (PWTs; 50% withdrawal threshold) to von Frey filaments, according to a modification of the method described by Chaplan et al.²³  Briefly, Semmes–Weinstein monofilaments (Stoelting Co., Wood Dale, IL) were applied to the plantar surface of the hind paw for 10 s or until the rat exhibited a flexion reflex. Filaments were applied in ascending (following no response) or descending (following a response) order of strength to determine the filament closest to the threshold of response. Filaments were applied thus, until a pattern of responses showing three changes (rats responding or not) was obtained.²⁴  The minimal stimulus intensity was 0.25 g, and the maximum was 15 g.

In order to rule out any sedative effects or motor disturbances elicited by SN50, rats were tested by being placed on a rotating drum (7 cm diameter), 23 cm above the floor of a metal enclosure (11 cm wide, 42 cm long, and 40 cm high) (Rotarod, IITC; Woodland Hills, CA). Normal, untreated rats were trained for 5 consecutive days, in which they underwent six trials, each separated by a 20-min recovery period. Each trial began by placing the rat on the stationary cylinder and then linearly increasing cylinder rotation from 0 to 30 rpm during the trial duration of 2 min. Time was measured until the rat fell from the rotating rod or till the end of the trial (latency to fall). On day 6, performance was recorded on the rotarod starting 30 min after injection of 600 ng of intrathecal SN50, using the experimental paradigm described above. Reported rotarod latency scores were based on the average of the six trials performed on each test day.

Glycemic values (mm of glucose), PWT (g), and ELISA results are expressed as mean ± 0.95 CIs. The choice of the appropriate analysis was based on sample size, variance homogeneity (Levene test), and on data distribution (Kolmogorov–Smirnov test of normality). Statistical differences among the CPIP groups were assessed using two-way ANOVA for one repeated measurement (groups by time during I/R). Analysis of PWT measurements from rats used for NFκB measurements was similarly performed by two-way repeated-measures ANOVA. Analysis of PWTs obtained after drug treatment was performed by three-way ANOVA with two repeated measures (treatment group by drug dose by time after drug administration). If significant heterogeneity of variance was detected (Levene test, P < 0.05), the ANOVA was performed using square root–transformed observations (raw, untransformed data are graphically presented). When appropriate, Tukey tests were used for post hoc pair-wise comparisons between groups. Within group, comparisons of test values to the corresponding baseline scores were also done using Dunnett test. An analysis of covariance was conducted on the linear regression models (area under the curve vs. log dose of SN50), and a Student t test (two-tailed) was conducted on the linear regression intercepts to assess shifts in the dose–response curve. Daily rotarod performance was analyzed using one-way repeated measures ANOVA followed by Tukey post hoc comparisons.

The test used and the degrees of freedom are indicated with each test statistic in the Results section. Results were considered significant if P value was less than 0.05. Statistics were performed using SPSS (Statistical Package for Social Sciences, 2004; IBM, Chicago, IL) or Statistica (Version 6; StatSoft, Tulsa, OK) software.

As shown in figure 1A, the four different groups of CPIP rats that underwent tissue extraction on day 2 postprocedure displayed statistically significant differences in glycemia during the I/R injury (F(18, 120) = 13.909, P < 0.0001, ANOVA group × time interaction). Although the mean glycemia of the four groups did not differ at baseline (P > 0.05, Tukey tests), insulin administration significantly lowered the blood glucose of the fed/insulin group below baseline values throughout the ischemic period at 30 min (P < 0.001), 90 min (P < 0.001), 150 min (P < 0.001), and throughout the period from 20, 40, and 60 min after reperfusion (P < 0.01 for each, Tukey tests). The glycemic level of the fed/insulin/dextrose group was less than baseline values only at 30 min postischemia (P < 0.05, Dunnett test). In contrast, dextrose administration to fed animals increased glucose levels compared with baseline at 90 min postischemia (P < 0.05). Rats in the fed group did not exhibit significant variations in mean glycemia at any time point. Compared with the fed group, rats of the fed/insulin group had lower glucose at 30 min of ischemia (P = 0.0343, Tukey tests), but not thereafter. Rats in the fed/dextrose group had significantly higher glucose levels compared with the fed group at 90 min of ischemia (P = 0.0318). Rats in the fed/insulin/dextrose group had higher glycemic values at 90 and 150 min of ischemia when compared with rats in the fed/insulin group (P = 0.0037 and P = 0.0017, respectively). Rats in the fed/insulin/dextrose group did not differ from rats in the fed group at any time point (Tukey tests).

The results of the PWT testing 48 h after the ischemic challenge are displayed in figure 1B. The PWTs varied significantly depending on glycemic treatment and injury (sham vs. CPIP; group by treatment interaction: F(3, 46) = 4.0943, P = 0.0117, ANOVA). In the fed, fed/insulin/dextrose, and fed/dextrose groups, CPIP animals displayed lower PWT compared with sham-treated rats (Tukey test; P = 0.0263, P < 0.0001, and P < 0.001, respectively). There was no difference in PWT between CPIP and sham-treated animals in the fed/insulin group. The CPIP animals in the fed/insulin group exhibited significantly higher PWTs compared with those in the fed group (P = 0.0049). The fed/insulin CPIP rats also exhibited significantly higher PWTs in comparison to CPIP rats in the fed/insulin/dextrose and fed/dextrose groups (P = 0.003 and P < 0.0001, respectively). There was no significant difference in the PWTs between the fed and the fed/insulin/dextrose groups, but PWTs in the fed/dextrose group were significantly lower than CPIP rats in the fed group (P = 0.0009). PWTs did not differ between groups for sham-treated rats. The relationship between mean glucose level measured during the ischemia period and each animal’s PWT is displayed in figure 1C. As shown, PWTs were negatively related to glucose levels, with a highly significant R² value.

As shown in figure 2A, NFκB levels in the ipsilateral hind paw muscles varied significantly depending on glycemic treatment (F(3, 42) = 7.633, P = 0.0004) and injury condition (F(1, 42) = 95.332, P < 0.0001). NFκB P65 levels were significantly higher in CPIP animals compared with sham-treated rats in the fed, fed/insulin/dextrose, and fed/dextrose groups (Tukey test; P = 0.0004, P = 0.0003, and P = 0.0002, respectively). CPIP rats of the fed/insulin group did not differ from the sham-treated control rats. The CPIP animals of the fed, fed/insulin/dextrose, and fed/dextrose groups also exhibited significantly higher NFκB levels in the plantar muscles compared with those of the fed/insulin group (P = 0.038; P = 0.024; P = 0.001).

As illustrated in figure 2B, dorsal horn NFκB levels were significantly different between the groups (F(1, 42) = 28.267, P < 0.0001). NFκB P65 levels were significantly higher in the ipsilateral dorsal horn of CPIP compared with sham-treated rats in the fed, fed/insulin/dextrose, and fed/dextrose groups (P = 0.0441, P = 0.0068, and P = 0.0080, respectively). CPIP rats of the fed/insulin rats did not differ from the sham-treated control animals. CPIP rats of the fed/dextrose group had higher P65 levels than CPIP rats of the fed/insulin group (P = 0.0246). No significant difference was found between the fed rats and the fed/insulin rats or between the fed rats and the fed/insulin/dextrose rats.

As depicted in figure 3A, the four groups of CPIP rats that underwent treatment with SN50 had varying mean glycemia during the ischemia and the subsequent reperfusion period (Group by time interaction: F(18, 150) = 8.38, P < 0.00001). The mean group glucose values did not differ at baseline. Glycemia in the fed group remained constant throughout the ischemia and reperfusion periods when compared with their baseline values (Dunnett test, P > 0.05). Glycemia was significantly reduced compared with baseline in fed/insulin rats at every time point during ischemia injury and reperfusion (P < 0.001, Dunnett test), and in the fed/insulin/dextrose rats only at 30-min postischemia (P < 0.05, Dunnett test). Conversely, glycemia was increased compared with baseline in fed/dextrose rats at 90 min into the ischemic period (P = 0.0002). Glycemic levels of the fed/insulin/dextrose rats were significantly higher than those of the fed rats at 20 min after reperfusion (P = 0.0242, Tukey test). Glycemia in the fed/insulin rats was significantly lower than that of the fed/dextrose group after 90 and 150 min of ischemia (P = 0.0021 and P = 0.0251, respectively, Tukey tests), and significantly lower than that of the fed/insulin/dextrose group also at 90 and 150 min of ischemia (P < 0.0003 and P = 0.0009, respectively), and at 30 and 40 min of the reperfusion period (P = 0.0001, P = 0.0132, respectively, Tukey tests).

As illustrated in figure 3B, significant differences were observed in PWT scores between the four different glycemic conditions at predrug baseline 2 days after I/R injury (F(1, 41) = 81.614, P < 0.0001). PWTs were lower in CPIP animals than in the corresponding sham-treated rats in the fed, fed/insulin/dextrose, and fed/dextrose groups (P = 0.0179, P = 0.0002, and P = 0.0002, respectively). There was no difference in PWTs between CPIP and sham-treated animals in the fed/insulin group). The fed/insulin group exhibited significantly higher PWTs than the fed/insulin/dextrose and the fed/dextrose groups (P = 0.0161 and P = 0.0031, respectively). However, there was no significant difference in PWTs between the fed and the fed/insulin/dextrose group (P > 0.05). The relationship between mean glucose level measured during the ischemia period and each animal’s PWT is displayed in figure 3C. As shown, once again PWTs were negatively related to glucose levels, with a highly significant R² value.

As stated in the Methods section, the CPIP animals of each glycemic treatment group received four different doses of SN50 and a single dose of an inactive control peptide corresponding to the scrambled sequence of SN50. A common dose of 200 ng SN50 was administered to each group. The PWTs of the fed/insulin group (fig. 4A) varied significantly as a function of the time and SN50 dose in a dose-dependent manner (ANOVA dose × time interaction, F(20, 120) = 1.66, P = 0.050). PWTs in this group did not change significantly over the course of testing after the administration of control peptide or 6.66 ng of SN50. After the administration of 22.22 ng of SN50, the PWTs of the fed/insulin group were significantly higher than baseline values at 30 and 75 min (P = 0.011 and P < 0.0001, Dunnett test). With 66.66 ng SN50, PWTs were significantly increased compared with baseline at 30 and 75 min postinjection (P = 0.012 and P = 0.003, respectively, Dunnett test). Finally, administration of 200 ng SN50 to the fed/insulin group led to significant increases in PWTs above baseline throughout the testing period at 30 (P = 0.012), 75 (P = 0.010), 120 (P = 0.007), 165 (P = 0.006), and 210 min postinjection (P = 0.006, Dunnett tests).

The PWTs of the fed group (fig. 4B) also varied as a function of time and SN50 dose in a dose-dependent manner (ANOVA dose × time interaction; F(20, 140) = 2.760, P = 0.0003). PWTs in this group did not significantly change at any point postdrug compared to the baseline levels after control peptide, 22.22 or 66.66 ng of SN50. Administration of 200 ng of SN50 resulted in a significant increase of the PWTs compared with baseline at 30 and 75 min postdrug administration (P < 0.0001 and P < 0.0001, respectively, Dunnett test). Finally, at 600 ng SN50, the PWTs were significantly higher than the baseline values for the whole duration of the testing period, at 30 (P = 0.004), 65 (P < 0.0001), 120 (P = 0.001), 165 (P < 0.0001), and 210 min postinjection (P < 0.0001, Dunnett test).

Rats of the fed/insulin/dextrose group (fig. 4C) also displayed dose-dependent effects of SN50 (ANOVA dose × time interaction; F(20, 120) = 3.614, P < 0.0001). Administration of 100 or 200 ng of SN50 did not significantly change the PWTs at any time point. SN50 400 ng significantly raised the PWTs of the fed/insulin/dextrose rats above the baseline values at 30 (P < 0.0001) and 75 min postinjection (P < 0.0001, Dunnett tests), whereas 600 ng of SN50 raised PWTs significantly at 30 (P <0.0001), 75 (P < 0.0001), 120 (P < 0.0001), and 210 min postinjection (P < 0.0001, Dunnett tests).

Rats of the fed/dextrose group (fig. 4D) displayed the lowest responsiveness to SN50 in the dose–response profile over time (F(20,120) = 2.1743, P = 0.0053). Administration of 200 ng of SN50 had no significant effect on PWTs, whereas 400 ng of SN50 only increased PWTs at 75 min postinjection (P = 0.0086, Dunnett test). The PWTs were increased above predrug baseline values at 75- and 120-min postinjection (P = 0.0065 and P = 0.0135, respectively) after 800 ng SN50. After administration of 1,000 ng SN50, PWTs were significantly increased above baseline values for every time point from 30 to 210 min postinjection (P = 0.0190, P = 0.0002, P = 0.0004, P = 0.0003, and P = 0.0042, respectively).

Figure 5A illustrates that the four CPIP groups reacted differently to the common 200 ng dose of SN50 (F(10, 90) = 2.380, P = 0.015, ANOVA group × time interaction). The fed/insulin group exhibited significantly higher mechanical thresholds compared with the fed/insulin/dextrose group at 30 (P = 0.013), 165 (P = 0.010), and 210 min postinjection (P = 0.007, Tukey tests), and compared with the fed group at 210 min postinjection (P = 0.035, Tukey test). The fed group exhibited significantly higher mechanical thresholds compared with the fed/insulin/dextrose group only at 30 min postinjection (P = 0.026, Tukey test).

Each group exhibited consistent baseline (predrug) PWTs, so there was no residual effect of SN50 before the administration of the next dose or any other interaction between day of testing and mean group PWTs (two-way ANOVA, F(12, 96) = 0.6216, P > 0.05, and F(4, 96) = 5.243, P > 0.05 for the days by treatment group interaction and days effects, respectively).

As illustrated in figure 5B, administration of different SN50 doses to the four groups produced log dose–response curves with slopes that did not differ significantly (F(1) = 5.05, P > 0.05, analysis of covariance, but with statistically significant shifts relative to each other (F(2) = 7.51, P < 0.01, analysis of covariance). The log dose–response curve of the fed/insulin/dextrose was shifted significantly to the right relative to the fed/insulin group, with a significantly lower x-intercept for the later (4.06 ± 7.04 vs. 77.80 ± 44.50 ng; P = 0.0262). The x-intercept of the log dose–response curve from the fed/insulin/dextrose group did not differ from that of the fed group (77.80 ± 44.50 vs. 24.89 ± 17.20, respectively; P = 0.088), but was higher than that of the fed/insulin group (P = 0.0345). The x-intercept for the log dose–response curve from the rats in the fed/dextrose condition (328.94 ± 92.94 ng) was also significantly higher than that of the fed/insulin groups (P = 0.031). The x-intercept value from the fed and fed/insulin did not differ from each other.

As illustrated in figure 5C, the rats’ rotarod performance improved throughout the 5-day training period (F(5, 210) = 2.661, P = 0.023, ANOVA). After the administration of 600 ng SN50 on day 6, rats stayed on the rotarod for an average of 65.14 ± 5.67 s, which was not significantly different from the performance achieved on day 5 (54.82 ± 5.22).

The current study has replicated our earlier study showing that varying glycemic levels at the time of I/R injury are associated with different postinjury mechanical allodynia. The current study further demonstrates that varying glycemic levels at the time of I/R is associated with varying NFκB levels in the ipsilateral hind paw muscles and ipsilateral dorsal horn of the spinal cord at 2 days post-IR injury; NFκB levels being higher in relatively hyperglycemic groups. These results suggest that NFκB plays a role in the pathophysiology of post-I/R mechanical allodynia in CPIP rats. Accordingly, administration of SN50 resulted in a reduction of postlesion mechanical allodynia in a manner associated with the glycemia present during the I/R injury, with the dose–response curve of the fed/insulin group being significantly shifted to the left in comparison with the fed/insulin/dextrose, fed, and fed/dextrose groups.

The four treatment groups included in the study differed in terms of glycemic values during I/R injury. Different glycemic values during I/R injury led to significantly different NFκB levels, in both the ipsilateral hind paw and the ipsilateral dorsal horn, suggesting that glycemic level affected not only peripherally activated NFκB, but also that glycemia influenced the central activation of NFκB, presumably induced by the peripheral I/R injury. Thus, tight glycemic control at the time of the injury seems to result in reduced NFκB levels peripherally and spinally, whereas, higher glycemia led to higher NFκB levels. These changes in NFκB levels were present 2 days post-I/R injury in both muscle and spinal cord.

The lower activated NFκB levels in the fed/insulin group are likely secondary to a lower glycemic level more so than antiinflammatory effects of insulin, as insulin administration to the fed/insulin/dextrose group did not prevent the increase in NFκB levels. This was also observed in a previous study examining the effects of decreasing or increasing glycemia during I/R injury on CPIP.⁴  Consequently, a lower glycemia per se seems more protective than other direct insulin effects in terms of NFκB levels. Importantly, the same relationship between these two groups was exhibited in the measures of mechanical allodynia, where insulin’s antiallodynic effects were lost when dextrose was added during I/R injury. These results are consistent with our previously published results showing that allodynia in CPIP rats decreases as much by fasting as by insulin treatment, and that giving dextrose to fasting CPIP rats eliminated the benefit produced by fasting.⁴  We predict, therefore, that fasting would also reduce NFκB levels in CPIP rats.

As we saw previously, different mean glycemic values between the fed, the fed/insulin the fed/insulin/dextrose, and the fed/dextrose groups during I/R injury led to significantly different postischemia allodynia, with a protective effect of lower glycemia on mechanical allodynia. Because higher glycemic levels are correlated with higher NFκB levels, NFκB may be the mechanism by which high glycemia worsens postischemic pain in the CPIP rat model. Indeed, activated NFκB has been demonstrated to play a role in several pain models with different etiologies, such as inflammatory,²⁵  neuropathic,^26,27  or postischemic²⁸  pain models. NFκB is activated by proinflammatory cytokines in the spinal cord.^29,30  Activated NFκB induces the transcription of inflammatory genes,³¹  and plays a key role in signaling pathways involving pain mediators, such as cyclooxygenase-2, dynorphin, c-fos, and neuronal nitric oxide synthase.^32–35  NFκB thus perpetuates spinal cord inflammatory responses, which further facilitate pain transmission, central sensitization, and chronic pain states.³⁶ 

The different response profiles of each experimental group to SN50 also points toward an important role of NFκB in the pathophysiology of CPIP, and the influence of glycemia on CPIP. The fed/insulin group (leftward shift) achieved significantly higher von Frey thresholds at a lower SN50 dose than the fed, fed/insulin/dextrose, or fed/dextrose groups. Furthermore, this effect was maintained for a longer period. This shows that rats with lower glycemia and NFκB levels require lower doses of SN50. This suggests that less inhibition of NFκB translocation is needed to alleviate CPIP. The dose–response findings are in accordance with a role of NFκB in allodynia induced by CPIP.²⁸  Furthermore, the varying sensitivity, and duration of responses to SN50 in the various groups implies that glycemia-related increased mechanical allodynia depends on NFκB activity.

In the current study, SN50’s effects on mechanical allodynia were maintained up to day 8 post-I/R. This finding is important because a previous study showed that NFκB levels in CPIP rats were normalized in muscle 7 days after the I/R, but were still increased in spinal cord tissue, suggesting that the central pathology was still ongoing, and that its inhibition can still interrupt the inflammatory cascade involved in nociception.²⁸  Although, we do not feel it is necessary to reconfirm the action of SN50 as a NFκB inhibitor, it would be useful to determine the relationship between allodynia and NFκB levels for each glycemic condition at later time points.

One may question the specificity of SN50, as at high concentrations it has been shown to block the nuclear import of not only NFκB, but also of other transcription factors such as activator protein 1, signal transducers and activator of transcription, and nuclear factor of activated T-cells³⁷  However, these observations occurred in cell lines and may not be transposable to in vivo experiments. Furthermore, the doses we used in our experiments were also significantly less than the 210 μg/ml used in the in vitro study.

In conclusion, we demonstrated that the differences in mechanical allodynia thresholds between the three groups of CPIP rats with varying glycemic levels are correlated with different NFκB levels present in the ipsilateral paw muscles and the ipsilateral dorsal horn of these rats, and that these different NFκB levels are modulated by the glycemia present during the I/R injury. Furthermore, the antiallodynic dose–response effects of SN50 were shifted to the left in relatively hypoglycemic rats, and to the right in relatively hyperglycemic rats. These observations could have important applications for postischemic pain conditions: by maintaining a tight glycemic control during I/R injury, less NFκB will be activated, and an analgesic response to an NFκB inhibitor will be attainable with smaller doses, and for longer periods of time.

The authors thank Réjean Huot, M.A., Retired Professor, Department of Psychology, Limoilou College and Laval University, Quebec City, Quebec, Canada, for his help with the statistical analysis of the results; Philippe Huot, M.D., M.Sc., F.R.C.P.(C.), D.A.B.P.N., Postdoctoral Fellow, Toronto Western Research Institute, University Health Network, Toronto, Ontario, Canada, for reviewing the article; and Karen Brown, M.D., F.R.C.P.(C.), Professor, Department of Anesthesia, Montreal Children’s Hospital, McGill University Health Center, Montreal, Quebec, Canada, for her useful suggestions.