Induction of thermal hyperalgesia and synaptic long-term potentiation in the spinal cord lamina I by TNF-α and IL-1β is mediated by glial cells

Abstract

Long-term potentiation (LTP) of synaptic strength in nociceptive pathways is a cellular model of hyperalgesia. The emerging literature suggests a role for cytokines released by spinal glial cells for both LTP and hyperalgesia. However, the underlying mechanisms are still not fully understood. In rat lumbar spinal cord slices, we now demonstrate that conditioning high-frequency stimulation of primary afferents activated spinal microglia within <30 min and spinal astrocytes within ~2 s. Activation of spinal glia was indispensible for LTP induction at C-fiber synapses with spinal lamina I neurons. The cytokines interleukin-1β (IL-1β) and tumor necrosis factor-α (TNF-α), which are both released by activated glial cells, were individually sufficient and necessary for LTP induction via redundant pathways. They differentially amplified 2-amino-3-(3-hydroxy-5-methyl-isoxazol-4-yl)-propanoic acid receptor-mediated and N-methyl-D-aspartic acid receptor-mediated synaptic currents in lamina I neurons. Unexpectedly, the synaptic effects by IL-1β and TNF-α were not mediated directly via activation of neuronal cytokine receptors, but rather, indirectly via IL-1 receptors and TNF receptors being expressed on glial cells in superficial spinal dorsal horn. Bath application of IL-1β or TNF-α led to the release profiles of pro-inflammatory and anti-inflammatory cytokines, chemokines, and growth factors, which overlapped only partially. Heat hyperalgesia induced by spinal application of either IL-1β or TNF-α in naive animals also required activation of spinal glial cells. These results reveal a novel, decisive role of spinal glial cells for the synaptic effects of IL-1β and TNF-α and for some forms of hyperalgesia.

Figure 1.

HFS activates microglia and astrocytes. Representative experiments show the changes in microglial (A–C) and astrocytic (D–F) specific markers after HFS of primary afferent fibers. A, D, Immunoreactivity for the microglia-specific marker Iba1 (A, n = 15 slices) and the astrocyte-specific marker GFAP (D, n = 16 slices) in untreated slices. B, 30 min after HFS, Iba1 immunofluorescence is increased on the ipsilateral site (right) but not contralaterally to HFS (n = 14 slices). C, 120 min after HFS, Iba1 levels are back to control again (n = 14 slices). E, Thirty minutes after HFS, GFAP immunoreactivity is unchanged compared with control (n = 16 slices). F, Two hours after HFS of primary afferents, GFAP shows increased immunoreactivity on the ipsilateral (right) and contralateral (left) side (n = 16 slices). G, Quantification of Iba1 and GFAP intensity 30 and 120 min after HFS. *p < 0.05; **p < 0.001; one-way ANOVA.

Figure 2.

Astrocytes respond to HFS with an intracellular Ca²⁺ rise. A, Example of a spinal cord slice loaded with the Ca²⁺-sensitive dye Oregon Green BAPTA 1-AM (OGB, green) and the astrocytic marker SR101 (red). Neurons are depicted with an arrowhead, astrocytes with an arrow. B, Time course of the HFS-mediated Ca²⁺ response of the three neurons (blue) and two astrocytes (purple) depicted in A. The Ca²⁺ rise in neurons is followed by a prolonged Ca²⁺ increase in astrocytes. C, Description of differences in Ca²⁺ kinetics between neurons and astrocytes. The increase in fluorescence and decay time was calculated between all neurons and all astrocytes recorded. ***p < 0.001.

Figure 3.

Spinal glia are necessary for LTP induction in vitro. Recordings were performed in lamina I neurons of the spinal cord dorsal horn. A, Example of HFS-induced LTP in one spinal lamina I neuron. Amplitudes of individual EPSCs were normalized to pretreatment values (dotted line) and plotted against time. The time point at which HFS was applied is depicted with an arrow. Insets show individual EPSC traces recorded at the indicated time points. B, Average time course of EPSC amplitude (mean ± SEM) of 19 lamina I neurons in which HFS was applied. In 13 of 19 neurons tested, HFS induces a significant increase in EPSC amplitude 30 min after conditioning stimulation (p < 0.001, n = 13 neurons, one-way RM ANOVA). C, Application of fluorocitrate (0.01 mm) to spinal cord slices inhibits LTP induction by HFS (n = 9 neurons; p < 0.05 compared with control conditions; Fisher's exact test). D, Averaged time course of nine neurons in which minocycline prevents the induction of LTP after HFS. Slices were incubated with minocycline (100 μm) for 1 h before the experiments and minocycline (20 μm) was bath applied during the entire experiment. C-fiber-evoked EPSC amplitudes stay at control values (dotted line) after HFS (30 min after HFS, p > 0.05, one-way RM ANOVA).

Figure 4.

IL-1β and TNF-α are necessary for LTP induction in vitro. Recordings were performed in lamina I neurons of the spinal cord dorsal horn. A, Bath application of IL-1ra (40 ng/ml) does not prevent LTP induction in 8 of 14 neurons tested (p > 0.05 compared with untreated slices, Fisher's exact test). B, A soluble TNF receptor (sTNFR1, 0.5 μg/ml) also does not inhibit LTP induction by HFS in 8 of 13 neurons recorded (p > 0.05 compared with untreated slices, Fisher's exact test). C, Application of both IL-1ra and sTNFR1 prevents LTP induction in all neurons tested (n = 9, p < 0.05 compared with untreated slices, Fisher's exact test).

Figure 5.

Potentiation of synaptic transmission by TNF-α and IL-1β. Patch-clamp recordings of spinal lamina I neurons show an increase in evoked excitatory synaptic transmission after bath application of the pro-inflammatory cytokines IL-1β and TNF-α. C-fiber-evoked EPSC amplitude (% control) is plotted against time (min). A, In 15 neurons tested, C-fiber-evoked EPSC amplitude stays constant over a recording period of 30 min (p > 0.05, one-way RM ANOVA). B, Bath application of TNF-α (20 ng/ml, 20 min) significantly augments C-fiber-evoked EPSC amplitude in 6 of 15 neurons tested (p < 0.05 compared with control, Fisher's exact test). C, Application of IL-1β (20 ng/ml) for 20 min increases synaptic transmission in 11 of 15 neurons tested (p < 0.001, compared with control, Fisher's exact test).

Figure 6.

Effects of IL-1β and TNF-α on synaptic transmission mediated by NMDA or AMPARs. Means ± SEM of NMDAR-evoked (A–C) or AMPAR-evoked (D–F) EPSC amplitudes were calculated and plotted against time (min). A, NMDAR-mediated currents do not change significantly over a recording period of 40 min (n = 5, p > 0.05, one-way RM ANOVA). B, Bath application of TNF-α increases synaptic transmission mediated by NMDARs in a subset (8 of 17) of lamina I neurons (p < 0.001, one-way RM ANOVA). C, In nine neurons recorded, IL-1β significantly potentiates NMDAR-mediated EPSC amplitude (p < 0.001, 20 min after wash-in of IL-1β compared with baseline, one-way RM ANOVA). D, AMPAR-mediated currents stayed stable over a recording period of 30 min (n = 5 neurons, p > 0.05, one-way RM ANOVA). E, TNF-α (20 ng/ml) significantly enhances AMPAR-mediated currents in nine of 23 neurons tested (p < 0.05, 20 min after TNF-α application compared with baseline). F, In eight neurons tested, IL-1β (20 ng/ml) has no significant effect on AMPAR-mediated currents (p > 0.05, 20 min after IL-1β compared with pre-observation).

Figure 7.

Effects of TNF-α on AMPAR trafficking and IL-1β on NMDAR phosphorylation (pNR1). Representative immunoblots are shown on top, and mean relative protein levels are displayed in the bar graphs. Spinal cords were obtained from 20- to 23-d-old rats. Spinal cords were incubated with TNF-α (20 ng/ml) or IL-1β (20 ng/ml) for 20 min. Data are displayed as mean ± SEM. A, TNF-α decreased cytosolic GluR1 significantly (n = 15 spinal cords, p < 0.01, t test) but had no effect on the membrane fraction of GluR1 (n = 15 spinal cords, p > 0.05, t test). B, Western blot analysis of spinal cords treated with IL-1β and TNF-α revealed that IL-1β significantly increased NMDAR phosphorylation (n = 15 spinal cords, p < 0.05, t test), whereas TNF-α did not significantly alter NMDAR phosphorylation (n = 15 spinal cords, p > 0.05, t test).

Figure 8.

Receptors for the pro-inflammatory cytokines TNF-α and IL-1β are expressed by lamina I neurons and glial cells. Double immunostaining shows colocalization of TNFR1 (Aa,Ba, green) and TNFR2 (Ca,Da,Ea, green) and IL1R (Fa,Ga,Ha, green) with the neuronal marker NeuN (Ab,Cb,Fb, red), the astrocytic marker GFAP (Bb,Db,Gb, red), and the microglial marker Iba1 (Eb,Hb, red) in spinal cord lamina I. Overlays reveal that lamina I neurons express TNFR1, TNFR2, and IL-1R. TNFR1 and IL-1R are also expressed by GFAP-positive cells in the spinal dorsal horn. There is weak costaining for the individual receptors and the microglial marker Iba1.

Figure 9.

Effects of IL-1β and TNF-α are not mediated exclusively by neurons. Shown are means ± SEM of AMPAR-evoked (A,B) or NMDAR-evoked (C,D) EPSC amplitudes calculated and plotted against time (min). A, In the presence of fluorocitrate (5 mm), AMPAR-evoked EPSC amplitudes stay stable over a recording period of 30 min (n = 15 neurons, p > 0.05, one-way RM ANOVA). B, When slices are incubated with fluorocitrate, TNF-α (20 ng/ml) has no effect on AMPAR-mediated currents (n = 15 neurons, p > 0.05, one-way RM ANOVA). C, In the presence of fluorocitrate, NMDAR-mediated currents do not change significantly in 10 neurons over a recording period of 40 min (p > 0.05, one-way RM ANOVA). D, After incubation with fluorocitrate, a 20 min application of IL-1β (20 ng/ml) significantly decreases NMDAR-mediated currents (n = 11 neurons, p < 0.001, one-way RM ANOVA).

Figure 10.

Inhibition of glial cells and IL-1β and TNF-α together prevents LTP induction after HFS in vivo. In A–E, the area of the C-fiber-evoked field potential is plotted against time (min). Data are expressed as mean ± SEM. A, Schematic drawing of the recording setup for in vivo experiments. B, Averaged time course of C-fiber-evoked field potentials recorded from eight rats in which HFS induces LTP (p < 0.001, one-way RM ANOVA). Values were normalized to pretreatment values (dotted line) and plotted against time. C, Spinal superfusion with the glial inhibitor fluorocitrate (0.01 mm) abolished the induction of LTP by HFS (n = 6 animals, p > 0.05, one-way RM ANOVA). D, Intravenous infusion of minocycline (60 mg/kg) beginning 1 h before the start of the experiment (open horizontal bar) prevents LTP induction by HFS (n = 6 animals, p < 0.001 compared with HFS control animals 60 min after HFS, t test). E, Spinal superfusion at the recording segment with IL-1ra (400 ng/10 μl) does not influence the establishment of spinal LTP after HFS (n = 7 animals, p < 0.05, one-way RM ANOVA). F, Spinal superfusion with sTNFR does also not prevent the induction of spinal LTP after HFS (n = 6 animals, p < 0.05, one-way RM ANOVA). G, The combination of IL-1ra and sTNFR prevents LTP induction after HFS (n = 5 animals, p > 0.05, one-way RM ANOVA). H, The p-p38 MAPK blocker SB203580 (200 μm) does not have any effect on HFS-induced LTP (n = 6 animals, p < 0.05, one-way RM ANOVA). J, A higher dose of SB203580 (400 μm) significantly reduces HFS-induced LTP compared with control animals 60 min after HFS (B; n = 5 animals; p < 0.05, t test).

Figure 11.

IL-1β- and TNF-α-induced hyperalgesia are prevented by glial cell blockade. A, Intrathecal injection of TNF-α (20 ng in 10 μl of saline) induces a significant reduction in the withdrawal latency in response to a radiant heat source compared with baseline levels (n = 9 animals, p < 0.001, two-way RM ANOVA). Intrathecal injection of fluorocitrate (1 nmol in 10 μl of saline) 2 h before TNF-α fully prevents cytokine-induced hyperalgesia (n = 9 animals, p > 0.05, two-way RM ANOVA). B, Spinal injection of IL-1β (20 ng in 10 μl of saline) induces heat hyperalgesia in nine animals tested (p < 0.001, two-way RM ANOVA). Fluorocitrate inhibits IL-1β-induced hyperalgesia significantly (n = 9 animals, p > 0.05, two-way RM ANOVA).