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We investigated the role and mechanisms of inflammatory responses within the dorsal root ganglion (DRG) in the development of chemogenic pathological pain. DRG inflammation was induced by a single deposit of the immune activator zymosan in incomplete Freund’s adjuvant in the epidural space near the L5 DRG via a small hole drilled through the transverse process. After a single zymosan injection, rats developed bilateral mechanical hyperalgesia and allodynia which began by day 1 after surgery, peaked at days 3–7, and lasted up to 28 days. The number of macrophages in ipsilateral and contralateral DRGs increased significantly, lasting over 14 days. Robust glial activation was observed in inflamed ganglia. Cytokine profile analysis using a multiplexing protein array system showed that, in normal DRG, all but IL-5, IL-10 and GM-CSF were detectable with concentrations of up to 180 pg/mg protein. Local inflammatory irritation selectively increased IL-1β, IL-6, IL-18, MCP-1, and GRO/KC up to 17 fold, and decreased IL-2 and IL-12 (p70) up to 3 fold. Inflaming the DRG also remarkably increased the incidence of spontaneous activity of A- and C-fibers recorded in the dorsal root. Many of the spontaneously active A-fibers exhibited a short-bursting discharge pattern. Changes in cytokines and spontaneous activity correlated with the time course of pain behaviors, especially light stroke-evoked tactile allodynia. Finally, local inflammation induced extensive sprouting of sympathetic fibers, extending from vascular processes within the inflamed DRG. These results demonstrate the feasibility of inducing chronic localized inflammatory responses in the DRG in the absence of traumatic nerve damage, and highlight the possible contribution of several inflammatory cytokines/chemokines to the generation of spontaneous activity and development and persistence of chemogenic pathologic pain.
Inflammatory responses in the lumbar dorsal root ganglion (DRG) play key roles in the development of many pathological pain states including chemogenic low back pain and some forms of intractable neuropathic pain. In humans, acute herpes zoster and postherpetic neuralgia result from inflammation in the DRG and along its dermatome following infection or reactivation of the varicella-zoster virus (Schon et al., 1987, Morgan and King, 1998). In laboratory animals, behavioral radiculopathy and sciatica can occur when the DRGs or dorsal roots are inflamed by exposure to material from the nucleus pulposus (NP) (Gertzbein et al., 1977, Kawakami et al., 1996, Olmarker and Myers, 1998, Satoh et al., 1999, Kawaguchi et al., 2001), which is known to possess immunogenic and chemogenic capacities (Gertzbein et al., 1977, Satoh et al., 1999, Kawaguchi et al., 2001).
More recently, it has been recognized that inflammation may also play a role in neuropathic pain (nerve injury) models even though these models do not use inflammation as the primary mechanism of initial injury. For example, inflammatory responses have been demonstrated in DRGs with peripheral nerve injury (Lu and Richardson, 1993, Eckert et al., 1999, Hu and McLachlan, 2002) or injury to the DRG (Zhang et al., 1999). In peripheral nerve injury, macrophages are recruited from blood to the damaged nerve and DRG, where they contribute significantly to the removal of degenerating axons and myelin sheaths, and to regeneration (Beuche and Friede, 1984, Brown et al., 1991, Perry and Brown, 1992, Hu and McLachlan, 2002). In addition, the satellite glia surrounding sensory neuron somata proliferate (Barron et al., 1990, Gehrmann et al., 1991, Lu and Richardson, 1991) and become immunoreactive for glial fibrillary acidic protein (GFAP) (Woodham et al., 1989).
Certain inflammatory cytokines within DRG or injured nerves are known to be associated with pain behaviors (Wagner and Myers, 1996, Sommer et al., 1998a, Sommer et al., 1998b, Ignatowski et al., 1999, George et al., 2000, Svensson et al., 2005). These cytokines play an important role in the development and persistence of nerve injury-induced cutaneous hypersensitivity (DeLeo and Colburn, 1995, Schafers et al., 2003a, Schafers et al., 2003b, Ji and Strichartz, 2004, Watkins and Maier, 2005, Zelenka et al., 2005), and also contribute to the generation of abnormal spontaneous activity from injured nerve fibers (Wagner and Myers, 1996, Sommer et al., 1998a, Sommer et al., 1998b, Ignatowski et al., 1999, George et al., 2000) or compressed DRG neurons (Liu et al., 2002, Zhang et al., 2002, White et al., 2005).
Little is known about how inflammation per se, in the absence of nerve damage, affects the DRG neurons, the cytokine profile within the DRG, and the subsequent animal pain behaviors. Numerous studies have examined inflammation or immune responses as the cause of low back pain/radiculopathy. However, most studies are limited to the acute phase due to the lack of an animal model of chronic epidural or DRG inflammation. Here, using a newly developed animal model of localized inflammation of the DRG (LID), we have examined possible causal factors that mediate pain following DRG inflammation. We characterized changes in cytokine production and their association with development of spontaneous activity as well as with pain and hyperalgesia. This study demonstrates the feasibility of the LID model in addressing neurological mechanisms of low back pain/radiculopathy and other pathologic states that involve inflammatory processes in the DRG.
Adult, male, Sprague-Dawley rats weighing 200–250 g were housed in groups of 2–4 in 40×60×30 cm plastic cages with soft bedding under a 12/12 h day/night cycle. The rats were kept 7–10 days under these conditions before surgery and up to 4 weeks after surgery. The experimental protocol was approved by the Institutional Animal Care and Use Committees of the University of Cincinnati and University of Arkansas for Medical Sciences.
Rats were anesthetized by continuous inhalation of isoflurane. On the right side, the paraspinal muscles were separated surgically from the L5-L6 vertebrae. The superior articular process of L6 and the transverse process of L5 vertebrae were cleaned. A small opening (diameter: 0.49 mm) was drilled through the junction of the transverse process and the lamina over the L5 DRG. A 25-G needle was cut short (length: 2–3 mm) and inserted into the opening without contacting the ganglion (Zhang et al., 2001). The needle was connected to a microsyringe loaded with 20 μl zymosan (Sigma) suspended in incomplete Freund’s adjuvant (IFA; Sigma) at a concentration of 0.5 μg/μl or 0.05 μg/μl corresponding to total amounts of 10 and 1μg, respectively. The microsyringe was left in place for 2–3 min after injection of the 20 μl zymosan/IFA to prevent solution leakage. Sham-operated rats received the same surgical procedure and injection of 20 μl of oxygenated artificial cerebrospinal fluid (ACSF) containing (in mM): NaCl 130, KCl 3.5, NaH2PO4 1.25, NaHCO3 24, Dextrose 10, MgCl2 1.2, CaCl2 1.2 (pH = 7.3). In experiments for cytokine detection and histology, both L4 and L5 DRGs were inflamed by individually depositing equal amount of zymosan (10 μg) in IFA in order to increase the amount of tissue obtained per animal.
In order to estimate the degree to which the injected zymosan would spread to contralateral and adjacent ganglia, we conducted experiments in 2 rats in which FluoSheres (Invitrogen) sulfate fluorescent microspheres were added to the zymosan/IFA injection solution. Equal volumes of microspheres and zymosan (10μg dose) were added to IFA. The diameter of the microspheres used was 4μm, chosen to be close to the average particle size for the zymosan (yeast cell wall) preparation as reported by the manufacturer (Sigma), and sulfate microspheres were used instead of more hydrophillic types because zymosan is relatively hydrophobic. Examination of whole ganglia mounts on POD5 indicated that fluorescence was distributed throughout the L5 DRG beneath the injection site. No dye spread to the contralateral DRGs was found. Also no dye was observed on the ipsilateral L4 DRG, although some beads appeared on the spinal nerve for the L4 DRG, possibly due to leakage along the region of spinal nerve where the L4 and L5 spinal nerves merge together.
Animals were inspected and tested every three days for 10 days prior to surgery, every three days for the first two postoperative weeks, twice during the 3rd week and twice at the end of 4 weeks postoperatively for a total of 12 testing sessions. For general observation, the rats were placed on a table and notes were made on the animal’s gait and the posture of each hindpaw and the conditions of the hindpaw nails and skin.
A total of 20 rats were randomly assigned to 3 groups: high dose zymosan/IFA (n = 7), low dose zymosan/IFA (n = 7), and sham control (n = 6). During the behavioral testing, the rat was placed in a clear plastic cage with a floor of plastic wire mesh. The cage was elevated so that stimuli could be applied to each hind foot from beneath the cage. Von Frey filaments with 100-μm cylindrical tips capable of exerting bending forces of 20, 40, 60, 80, 120 and 160 mN were applied to 6 designated loci on the plantar surface of the foot. The percentage of withdrawal responses was plotted as a function of force; the response threshold was defined as the force corresponding to a 50% withdrawal, as determined by the Hill Equation (Zhang et al., 2000, Homma et al., 2002). The baseline mean withdrawal threshold of each hind paw before surgery was obtained from an average of 3 testing sessions, 1 session every 3 days (plotted as day 0). For each postoperative testing session, difference scores were computed by subtracting the baseline withdrawal threshold from each postoperative day threshold. Negative scores indicate a decreased threshold after surgery.
A total of 19 rats, 12 zymosan-treated, and 7 sham control, were tested for foot withdrawal to innocuous tactile stimulation before and after surgery. A wisp of cotton pulled up from, but still attached to a cotton swab was stroked mediolaterally across the plantar surface of the skin through the floor of the plastic cage. Six strokes were delivered to each hind paw, alternating between right and left with each stroke. A single quick withdrawal reflex was considered to indicate the existence of tactile allodynia. The incidence of withdrawals was expressed as a percentage of the six strokes for each foot. The stimulus was defined as normally innocuous because control rats never exhibited a withdrawal response.
In preliminary experiments, sensitivity of the paw to a thermal heat stimulus was examined up to POD 17 using the method of Hargreaves et al. (Hargreaves et al., 1988). These experiments showed that zymosan/IFA treatment resulted in only very small increases in thermal sensitivity (n = 6). The largest changes were observed on day 11, when the withdrawal threshold fell from a baseline value of 9.6 ± 0.4 seconds to 8.2 ± 0.3 seconds (p = 0.1, Student’s t-test) on the ipsilateral side, and from 10.2 ± 0.9 seconds to 8.3 ± 0.2 seconds (p = 0.02, Student’s t-test) on the contralateral side. Repeated measures ANOVA did not reveal significant changes in thermal sensitivity from baseline on any of the postoperative days tested (up to POD 17). For comparison, we observed thermal sensitivity decreases of 3 to 5 seconds, from a similar baseline value, in our previous study using the CCI and SNI pain models (Xie et al., 2005). Because thermal pain sensitivity changes were small and nonsignificant, we focused on mechanical pain sensitivity for the rest of this study.
A 10-μm frozen section was cut from the harvested DRG every 30 μm, picked up on polylysine coated slides, and allowed to air-dry overnight. Sections were first incubated with rat-absorbed mouse anti-rat ED-2 (1:1,000) (Serotec, Inc., Raleigh, NC), a specific marker for macrophages, or rabbit anti-rat GFAP (1:800) (Immunostar, Hudson, WI), a marker for activated glia, overnight at room temperature followed by a reaction with horse anti-mouse and goat anti-rabbit biotinylated secondary antibodies, respectively (Vector Laboratories, Burlingame, CA). The immunoreactive products were visualized using the diaminobenzidine (DAB) method for ED-2 and DAB or Alexa Fluor 594 fluorescent dye (Invitrogen Corp., Carlsbad, CA) for GFAP.
Using ImagePro Plus software (Media Cybernetics, Inc., Silver Spring, MD), non-overlapping, adjacent images from each section were digitized under a light microscope equipped with a color digital camera and stored in a computer. To estimate the density of macrophages, positively stained macrophage profiles were counted and then normalized by the number of neurons in the analyzed image area to give a density ratio (number per neuron).
Cytokine expression profiles in the normal and inflamed DRG were evaluated using Bio-Plex System (Bio-Rad, Hercules, CA, USA) combined with Linco 14-Plex Rat Cytokine Detection Kit. A total of 14 rat cytokines were measured simultaneously from a single well according to the manufacturer’s protocols. Briefly, normal, sham, or inflamed ipsilateral DRGs (L4 and L5) were isolated from the rats on postoperative days (POD) 1, 3, 7, 14, 21, and 28. Tissue was homogenized (3 pulses, 15 sec each with a 3 min interval between pulses) using a PowerGen 125 at 4C° in lysis buffer (Bio-Rad, Hercules, San Diego, CA) supplemented with a protease inhibitor cocktail (Sigma, St. Louis, MO) followed by centrifugation (13,000 rpm) at 4°C for 30 min to obtain extracted protein.
The cytokine detection kit contains color-coded microspheres conjugated with a monoclonal antibody specific for a target protein. Protein samples (25 μl) extracted from DRG tissue were thawed and run in duplicate. Antibody-coupled beads were incubated with the tissue sample (antigen) after which they were incubated with biotinylated detection antibody before finally being incubated with streptavidinphycoerythrin. A broad sensitivity range of standards (Linco Research, Inc. MI, USA) ranging from 4.88 to 20,000 pg/ml was used to allow the quantization of a dynamic wide range of cytokine concentrations and provide the greatest sensitivity. This captured immunoassay was then read by the Bio-Plex System which uses Luminex fluorescent-bead-based technology (Luminex Corporation Austin, TX, USA) with a flow-based dual laser detector with real-time digital signal processing to detect both the color code and the signal intensity of each bead, allowing simultaneous quantification of multiple proteins in a single sample. This method was chosen because it has sensitivity and performance similar to ELISA methods, but requires much smaller sample volumes and is suitable for mutiplexing (duPont et al., 2005).
The concentrations of cytokines in these assays were calculated using a standard curve. A regression analysis was performed to derive an equation that was then used to predict the concentration of the unknown samples. The measured cytokines were interleukin-1α (IL-1α), IL-1β, IL-2, IL-4, IL-5, IL-6, IL-10, IL-12 (p70), IL-18, interferon-γ (IFN), tumor necrosis factor-α (TNF-α), granulocyte-macrophage colony stimulating factor (GM-CSF), growth related oncogene (GRO/KC) and monocyte chemoattractant protein-1 (MCP-1).
The final concentrations of the 14 cytokines were obtained from an average of values observed in 3 to 5 samples for each time point and condition. Each individual sample contained 4 DRGs combined from 2 rats, measured in duplicate. Data were normalized to the amount of protein and to the tissue weight, as indicated.
As described in our earlier publication (Zhang et al., 1997), under general anesthesia, a laminectomy was performed at the L3–L6 level, and the L5 DRG with attached sciatic nerve and dorsal root was mounted in a recording chamber after being removed from the rat. The DRG was secured by fine pins to the Sylgard base in the central chamber (5 mm in diameter). The dorsal root was led from this chamber and into an adjacent chamber (filled with mineral oil), where microfilament recordings were performed. The sciatic nerve was covered with Vaseline, and a bipolar-stimulating electrode was used for electrical stimulation. The DRG was continuously perfused at a rate of 5 ml/min with ACSF. The temperature was kept at 37° ± 0.5°C by running the perfusing solution through a temperature-controlled inline heater (Warner Instruments Corporation, Hamden, CT). Electrical activity was recorded extracellularly via laboratory interface and computer acquisition software (Spike2, Cambridge Electronic Design, Inc, England). Nerve fibers were classified as C- (<1.5 m/s), Aδ- (1.5–15 m/s), or Aβ-fibers (>15 m/s) based on the conduction velocity of impulses evoked by electrical stimulation of the nerve. The Govrin-Lippmann and Devor method (Govrin-Lippmann and Devor, 1978) was used to measure the incidence of ongoing discharge in those fibers where conduction velocity could be measured by electrical stimulation of the sciatic nerve. For each fiber bundle of approximately equal diameter (≈40–50 μm), the sciatic nerve was stimulated with a gradually increasing intensity of current (0.1–0.5 ms square wave pulses, 1–2 Hz) up to 10 mA, resulting in a gradual recruitment of dorsal root fibers until the number of fibers saturated. The total number of different action potential waveforms was counted and summed for all strands and divided by the total number of fibers (in all strands) recruited by electrical stimulation of the sciatic nerve to obtain the incidence of ectopic discharge.
Tissue sections (40 μm) were cut using a Vibratome (St. Louis, MO) and incubated in antibodies to tyrosine hydroxylase (TH) (raised from rabbit; obtained from Pel-Freeze, Rogers, AR) at a dilution of 1:1,000 for 48 hours at 4 °C, followed by reaction with biotinylated secondary antibody and, finally, with Vector ABC reagent. Triton-X (0.3%) was used in all reaction solutions to enhance antibody penetration. Immunoreaction products were visualized by the DAB method in the presence of H2O2 in 0.1 M phosphate buffer. Tissues were then mounted on gelatin-coated slides, air dried, dehydrated, and coverslipped for light-microscopic observation.
Data are expressed as means and standard errors of the mean. Differences in withdrawal thresholds over time were tested using one-way Repeated Measures Analysis of Variance (RM ANOVA) followed by all pairwise posthoc comparisons (Tukey method). Difference in withdrawal thresholds between groups (e.g., high dose vs. low dose) was tested using two-way RM ANOVA. Cytokine concentrations below the detection level of the experiment were assigned a value of zero for purposes of calculating average concentrations. Differences in macrophage densities between groups were compared using Student’s t-test. P<0.05 was chosen as the criterion for significance.
All animals appeared in good health throughout the testing period. The level of general activity was normal before and after surgery. Their fur was sleek and well groomed. No changes in gait or posture were observed in any rats of the sham or zymosan groups before or after surgery. There were no signs to indicate that rats were trying to reduce the weight placed upon the ipsilateral paw by leaning to the other side or by sitting on the opposite haunch as observed in other rodent models of pathological pain. However, when a mechanical or thermal stimulus was applied to the hind paws during postoperative testing, the reflex withdrawals were of great amplitude and excessive duration during which the paw was held in the air typically 2 – 15s. At times, a withdrawal of the hind paws was accompanied by exaggerated aversive behavior such as licking the stimulated paw. When the same stimuli were applied to the hind paw during preoperative testing, the reflex withdrawals were brief and of smaller amplitude.
Local application of zymosan/IFA induced bilateral hyperalgesia for up to 28 days post-operatively. Ipsilateral withdrawal thresholds dropped significantly on the first postoperative day for both high (10 μg, n=7) and low (1μg, n=7) doses of zymosan. Withdrawal thresholds steadily decreased until reaching a minimum threshold on POD 3 (p<0.05, One-way RM ANOVA). Threshold levels then began to steadily approach, but never returned to, pre-operative levels over the next 3 weeks. A higher dose of zymosan substantially decreased the withdrawal threshold when compared to the lower dose zymosan or the sham control (Figure 1A, B) (p<0.05, two-way RM ANOVA).
Local zymosan/IFA application on the DRG also induced an increased sensitivity in the contralateral foot, albeit with a slower onset. As was the case for the ipsilateral side, thresholds steadily approached, but never met, pre-operative levels through POD 28. Rats receiving buffered saline injection showed a slight, bilateral decrease in withdrawal threshold, which quickly returned to pre-surgical level before POD 7 (Figure 1C).
Prior to surgery, none of the 12 rats in the zymosan group (10 μg) exhibited any reflex withdrawals to light strokes of either foot with the cotton wisp. On the first postoperative day, however, light strokes of the ipsilateral or contralateral foot evoked a reflex withdrawal to at least one of the 6 strokes (tactile allodynia). The incidence of foot withdrawal to stroking was higher for the ipsilateral than the contralateral foot. The incidence of light stroke-evoked reflexes peaked on POD 3 and decreased during subsequent testing. By POD 14, none of the rats withdrew to the stimulus applied to either foot (Figure 1D). None of the 7 ACSF-treated sham rats exhibited any quick withdrawal response to light strokes with the cotton wisp during the entire 28-day testing period (data not shown).
Depositing zymosan (10 μg) in IFA on the L5 DRG resulted in prolonged macrophage response in the DRG (Figure 2). Ipsilaterally the macrophage density ratio (# of macrophage per neuron) peaked at POD 7, increasing to 0.78, nearly four times that of normal DRGs and more than triple that of the saline control (Figure 2, Figure 3, top panel). Density ratios remained considerably above normal while decreasing after POD7 with a POD 14 density ratio of 0.6; which was still nearly double the density of normal DRGs. Contralateral values increased more gradually with a POD 2 density ratio of 0.3 and a peak density ratio of 0.4 at POD 14 (Figure 3, bottom panel). Macrophage responses were also observed in DRG treated with low dose zymosan (1 μg). Macrophages were diffusely located throughout the inflamed DRG.
As shown in Figure 4, incubation of inflamed DRG sections, yielded strong rings (activated satellite glia) of signal around almost all of the large, medium and small-sized neurons that intensified with time within the first 2 postoperative weeks followed by a slow recovery but never returned completely to the presurgical level by 3 weeks postoperatively (Figure 4A, B). GFAP-positive rings were also observed in some areas of the contralateral DRG sections (Figure 4C). However, incubating normal DRG sections with the anti-GFAP antibody yielded little to no signal (Figure 4D).
As shown in Table 1, in normal DRG, of the 14 cytokines examined, all but IL-5, IL-10, and GM-CSF were detectable. Local zymosan/IFA application (10 μg) remarkably increased IL-1β, IL-6, IL-18, MCP-1, and GRO/KC up to 17 fold with much larger increases in zymosan-treated animals than in sham animals (Figure 5). Levels of IL-2 and IL-12 (p70), decreased by up to 3 fold although decreases of similar magnitude were also observed in sham operated animals (Figure 6). Levels of IL-1α, TNF-α and IFN were not strongly or consistently affected by localized inflammatory irritation. Levels of IL-4 seemed to decrease, in that a larger fraction of samples had concentrations below the detection level, but because the levels even in normal rats were close to the detection limit, quantification of decreased levels could not be obtained, and differences between sham and zymosan treated rats were not clearly evident.
As shown in figure 5, the largest changes in cytokine levels were observed on days 1 and 3. Most cytokines that had increased, declined back towards control levels considerably by day 7, though some still remained above control levels. An exception was IL-18, which showed similar increases through day 21. The two cytokines (IL-12 and IL-2) for which we were able to measure zymosan/IFA-induced decreases also showed changes that lasted through day 21.
Spontaneous (ectopic) activity, thought to be an important correlate of pathological pain in many chronic pain models (Wall et al., 1979, Burchiel, 1984, Devor et al., 1992, Xie et al., 2005), was examined in normal ganglia and those inflamed with zymosan (10 μg) in IFA. As summarized in Table 2, C-fibers were recorded from inflamed DRG via electrical stimulation of the sciatic nerve on POD 3, 7 and 14. Of the C-fibers, 4 to 11% were spontaneously active with the highest and only significantly different percentage observed on POD 3. All of the spontaneously active C-fibers recorded exhibited discharges with irregular interspike intervals and a low frequency that was typically 0.5 to 1 Hz. Spontaneous activity was also recorded from 5 to 9% of Aδ-fibers in inflamed ganglia, however, these increases did not reach statistical significance. The highest incidence of spontaneous activity was observed in Aβ-fibers, 10 to 16 % of which exhibited ectopic spontaneous activity within the inflamed DRG.
Thirty-two percent of the spontaneously active Aβ-fibers had a bursting pattern (Figure 7). The number of spikes within a single burst was 2 to 7 spikes per burst in the majority of these fibers (82% fell within this range on day 3, falling to 63% on day 7 (p<0.0001 vs. day 3), and to 59% on day 14), though a minority of fibers had much larger values of 10 – 40 spikes per burst, bringing the average number of spikes per burst up to 6.7 ± 1.3 on day 3. On days 7 and 14 a small number of fibers (6 – 12 %) displayed ultra long bursts of over 100 spikes. The interspike interval within each burst decreased from 39.6 ± 1.6 msec on day 3 to 26.8 ± 3.9 msec on day 14 (p < 0.01), with an intermediate value on day 7.
The patterns of the remaining spontaneously active, nonbursting fibers were either irregular or regular firing without bursting. Fibers with high-frequency (>15 Hz), irregular discharges were rare. The incidence of spontaneous activity was also measured for 4 unoperated normal rats. As previously observed in our lab (Zhang et al., 2002), the incidence of spontaneous activity was quite low (Table 2).
In 6 zymosan-treated rats (10 μg), the ipsilateral DRGs were dissected on POD 3 (n=2), 7 (n=2) or 14 (n=2) for immunohistochemical staining of the sympathetic fibers. In our previous studies, we have found that in normal DRGs, “dark cells” with TH-IR “tails” were scattered throughout the sections. TH-positive sympathetic fibers were rare. No abnormal growth of the sympathetic fibers was found in any sections from any vascular process (Zhang et al., 2004). In the inflamed DRGs, however, robust sprouting of the sympathetic fibers was observed in all sections beginning as early as day 3 after surgery. It was shown clearly in most sections that the sprouted fibers originated from the vascular processes (Figure 8A, B). The level of sprouting further increased on day 7 and persisted on day 14. As in axotomized DRGs, the sprouted fibers sometime formed basket-like structures around large- and medium-sized DRG neurons (Figure 8C, D).
Mechanical deformation and/or inflammatory irritation of the DRG and its nerve roots is a possible consequence of certain disorders, such as spinal stenosis, disk herniation, spinal injury, or tumors (Hoch et al., 1993, Sato and Kikuchi, 1993, Briggs and Chandraraj, 1995, Poletti, 1996). Clinical studies indicate that inflammation in the vicinity of the DRG may occur with or without apparent anatomic abnormalities. The mechanisms underlying inflammatory responses in the DRG and resultant low back pain and painful radiculopathy may be different in compressive vs. non-compressive injuries. In our previous studies, we have found that mechanical compression of the DRG by insertion of a metal rod into the intervertebral foramen induces pain and hyperalgesia by enhancing sensory neuron excitability (Song et al., 1997, Zhang et al., 1999) and possibly by causing synthesis and release of inflammatory cytokines within the compressed DRG (Homma et al., 2002). In the present study, we have created a new animal model, which will allow us to study inflammation, independently from compression or injury, as a biological factor contributing to the development of radiculopathy and sciatica. It was found that pain and hyperalgesia occurred in all rats after an injection of a small amount (1 to 10 μg) of the immune activator zymosan in IFA over one L5 DRG. The mechanical threshold to evoke a painful withdrawal response dropped to the lowest level within a week and remained low during the first two weeks. The tactile allodynia evidenced by painful foot withdrawal to light strokes using a piece of cotton wisp was present in all rats for the first two weeks. Although light stroke was not able to evoke withdrawal responses after POD14, cutaneous sensitivity remained higher than pre-surgical levels through the rest of our testing period. The time course of these pain behaviors correlated with the inflammatory responses in the DRGs as indicated by elevated numbers of macrophages and satellite glia between day 1 and day 14. In preliminary experiments, we found that inflaming the L5 DRG did not alter the withdrawal thresholds of the ipsilateral or contralateral hind paws to thermal stimulation. Thus, in the present study, we have focused on changes in mechanical sensitivity.
The rat cytokine protein array used in the present study can simultaneously detect 14 different cytokines with high specificity and sensitivity, comparable to that obtained with ELISA (duPont et al., 2005). This is the first protein level, cytokine profiling of the DRGs. It shows that in normal DRGs most cytokines are detectable. Zymosan/IFA treatment significantly altered cytokine levels. Consistent elevations, well above the levels seen in sham-operated animals, were observed for IL-1β, IL-6, IL-18, MCP-1, and GRO/KC, cyokines that are traditionally classified as pro-inflammatory based on their effects on classical immune cells; in contrast, anti-inflammatory cytokines IL-2 and IL-12 declined.
In the main these results are consistent with those seen in DRG in some other pain models. Pro-inflammatory cytokines are often elevated in pain states (as well as in other pathologies of the nervous system) and often have anti-analgesic roles though this can not be uniformly assumed (Watkins and Maier, 2002). However, these cytokines may also play protective roles in nervous system pathologies, and it is not yet clear how findings on the neuroprotective roles of pro-inflammatory cytokines in the CNS (Stoll et al., 2000, Felderhoff-Mueser et al., 2005) will apply to pathological pain, in which abnormal neuronal activity rather than neuronal death may play a more important role. Some of the elevated cytokines we observed, namely IL-1β, IL-6, and MCP-1, have previously been found in DRG in many studies; their receptors shown to be present on neurons and/or satellite glia; and their levels increased in various pain models. Several lines of evidence indicate the importance of these cytokines in mediating chronic pain states: i.t. and peripheral application of these cytokines can induce or enhance pain states; genetically modified mice lacking particular cytokines or their receptors have reduced pain in various models; and these cytokines have acute and long-term excitatory effects on DRG neurons (including nociceptors), and can enhance release of pain neurotransmitters (for review, see Watkins and Maier, 2002, Sommer and Kress, 2004).
One relatively new finding in our cytokine profile is the large increase in GRO/KC (also called CXCL1) in LID. There are few previous studies on the role of this cytokine in DRG, though it has been shown to enhance peptide release from neonatal DRG nociceptors in vitro and to induce hyperalgesia after peripheral injection (Qin et al., 2005). However, GRO/KC has been implicated in several CNS pathologies (e.g., Xia and Hyman, 2002, Losy et al., 2005, Valles et al., 2006), and can acutely modulate activity of cerebella neurons (Giovannelli et al., 1998). Similarly, there are few studies on the role of IL-18 in DRG, though it has been extensively studied in models of various CNS pathologies (Felderhoff-Mueser et al., 2005). In our study, IL-18 was notable for the more prolonged time course of its elevation.
Another interesting result of our cytokine measurements was that we found no consistent increases in TNF-α level in the inflamed DRG. In fact, the level on day 3, near the peak of pain behavior effects, we observed a small decrease (Table 1). This change was confirmed using a different cytokine assay kit from Bio-Rad (data not presented here). In our previous study (Homma et al., 2002), we found that perfusion of the compressed DRG with soluble TNF-α receptors led to a very small, albeit significant, decrease in pain behaviors. Thus, our data suggest that TNF-α plays only a minor role in the pain induced by the DRG compression or localized inflammation (e.g., LID). This contrasts with the extensive evidence for a role of TNF-α in peripheral nerve injury and peripheral inflammation models. These results highlight the importance of examining a number of different cytokines, now that this has become technically feasible.
The sources of cytokines in the inflamed DRG are not yet known. Activated satellite glia and macrophages are known to produce a number of cytokines in the early phases of inflammatory process and thus could contribute to the increased cytokine levels in the inflamed DRG. The slower time course of glial activation and macrophage infiltration compared with pain behaviors and changes in cytokine levels in the inflamed DRG might suggest that glia or macrophages are not likely to be the major sources of the elevated cytokines or a major cause of the pain behaviors. However, this interpretation is limited by the fact that these cells might rapidly release cytokines such as IL-1β, IL-18, IL-6, and TNF-α by converting them from the pre-existing pools of the inactive forms into the active, releasable forms – such a release might occur much more rapidly than upregulation of glial activation marker proteins that we measured, and in addition would presumably not be detected by our antibody-based detection system. Certain cytokines/chemokines such as MCP-1 and IL-1β can be synthesized and released from the sensory neurons in the DRG (Copray et al., 2001, White et al., 2005). Another possible source of chemotactic cytokines is the endothelial cells which can be activated by pro-inflammatory cytokines (Martiney et al., 1998).
The incidence of spontaneous activity in dorsal root fibers was high on POD 3–7 and dramatically decreased to a lower level by POD14. The time course for myelinated A-fibers (e.g., Aδ- and Aβ-) correlated well with the inflammatory response as well as the development of painful behaviors. In a recent in vitro study, it was found that topical DRG application of an inflammatory cocktail consisting of histamine, prostaglandins E2 (PGE2), serotonin (5-HT), and bradykinin robustly enhanced the excitability of the sensory neurons with intact or injured peripheral axons (Song et al., 2003). In another study, MCP-1 was found to excite chronically compressed DRG neurons in vitro (White et al., 2005). These results and results from our current study have led us to hypothesize that the initiation or development of abnormal spontaneous activity from inflamed DRG neurons may be attributed to the increased cytokine production and possibly to resultant elevated levels of inflammatory mediators within the DRG. The incidence of spontaneous activity with a bursting pattern is significantly higher shortly after surgery, especially the short-burst discharges. Similar observations were made in chronically compressed DRGs (Song et al., 1999). This suggests that in compressed DRGs, the bursting discharges may be caused by intraganglionic inflammation.
Sympathetic sprouting occurred in all inflamed DRGs beginning as early as day 3 of surgery. It can be clearly observed that sprouted fibers originate from the vascular processes in the DRGs. Fiber sprouting may be explained by increased expression of certain inflammatory cytokines such as IL-6, which has been reported to be able to cause sympathetic growth (Ramer et al., 1998, Ramer et al., 1999). Other factors such as NGF and NT-3 may be involved, too (Zhou et al., 1999). Results of this study indicate that sympathetic sprouting may occur in intact DRG without peripheral axotomy. Extensive sympathetic sprouting in the inflamed DRG suggests a possible sympathetic component in this model as reported in other inflammatory pain conditions (Nakamura and Ferreira, 1987).
It is of interest to compare the LID model with the previously described sciatic inflammatory neuritis (SIN) model of Chacur at al. (2001) in which pain was induced by injecting zymosan around the sciatic nerve. Both models produce mechanical but not thermal pain behaviors. SIN is associated with localized increases in IL-1β, as we observed in LID, but also results in increased TNF-α release. Both models produce both contralateral and ipsilateral pain at higher zymosan doses. Milligan et al. suggested that this phenomenon was possibly mediated by glial activation across the spinal cord. They found that the bilateral allodynia induced by SIN can be prevented or reversed by intrathecal application of glial inhibitors, inhibitors of p38 mitogen-activated kinases, or proinflammatory cytokine antagonists specific for interleukin-1, TNF-α, or interleukin-6 (Milligan et al., 2003). Contralateral allodynia and hyperalgesia in rats with zymosan/IFA treatment of a single DRG may share similar mechanisms. The possibility of zymosan diffusion in contributing to the development of contralateral allodynia cannot be completely ruled out, though our experiments with fluorescently labeled microspheres argue against this possibility.
Evidence for spinal contributions to pain induced by DRG inflammation was also obtained in another study, in which topical application of autologous NP to the L5 DRG remarkably increased the sensitivity of WDR neurons in response to mechanical or thermal stimulation of the peripheral receptive fields (Cuellara et al., 2004). Both NP and zymosan/IFA are considered to be immune activators and thus may share similar peripheral and spinal mechanisms in causing pain and hyperalgesia.
In summary, the current study demonstrated that localized immune activation in the vicinity of the L5 DRG induced prolonged inflammatory responses and elevated cytokine production along with enhanced spontaneous activity of DRG neurons and subsequent development of bilateral allodynia in the rats. These results have demonstrated a promising animal model for the study of neurological mechanisms of chemogenic low back pain, radiculopathy and sciatica, which may also become a reliable tool for the development and assessment of new therapies targeting these types of pathological conditions.
This work was supported in part by NIH grants NS39568 and NS45594 (J.Z.).