Chronic pain frequently coexists with other disorders such as anxiety, depression, or learning and memory deficits. This co-occurrence might manifest if chronic pain were to precipitate neuroplastic changes in supraspinal structures that lead to these pain-related disorders. The hippocampus is a key brain area recently implicated in the affective and cognitive impairment associated with chronic pain. Indeed, data from patients experiencing pain suggest that functional disturbances in frontohippocampal connectivity are a relevant cause of pain-related working memory deficits1,2; moreover, elderly patients who suffer from chronic pain appear to have a smaller hippocampus.3 Other studies in animal models of neuropathic pain have identified the hippocampal abnormalities associated with short-term4 and recognition memory deficits,5 as well as deficits in long-term potentiation.6 In addition, animals in which neuropathic pain is induced seem to be unable to extinguish contextual fear and show enhanced depressive- and anxiety-like behavior.7,8
One process that seems to mediate memory formation, as well as recognition of dangerous and stressful signals, is hippocampal neurogenesis.9–12 This process appears to occur in virtually all mammalian species, including humans,13,14 and we now know that the new cells generated in the hippocampus (neuroblasts) are incorporated into the granule cell layer, attaining the morphologic and biochemical characteristics of neurons.15,16 Thus, chronic pain might impair the generation of these new neurons, impeding appropriate learning and memory and thereby contributing to the development of mood disorders. Furthermore, recent data highlight a specific link between neurogenesis and neuropathic pain, with researchers showing that neuropathic pain decreases the number of neuroblasts8–10 and that it impairs the neurogenesis associated with enriched environments.17 The effects of pain on the survival and differentiation of newborn cells, however, still remain unknown, and the possible influence of stress on the neurogenic effects of pain is still to be demonstrated.
Stress consistently has been shown to be a factor that contributes to the maintenance and amplification of the severity of pain. However, most reports related to such issues are correlational, and the underlying biologic mechanisms remain unclear. However, the relevance of such findings are evident, given that clinical observations suggest that stress increases the susceptibility of developing pain and that it exacerbates existing pain.18–20 For these reasons, we set out to confirm the effects of neuropathic pain on cell proliferation and to investigate its possible effects on the survival and differentiation of cells generated de novo in the dentate gyrus (DG) of the rat hippocampus. In addition, we aimed to define the cellular consequences of adding moderate stress to the neurogenic effect of pain to mimic the daily stressful situations experienced by patients with chronic pain.
Experiments were performed on male Sprague-Dawley rats (150–180 g, 6–8 weeks old at the time of surgery) that were housed in standard conditions (22°C–24°C, relative humidity of 45%–65%, 12-hour light/dark cycle with ad libitum access to food and water), performing all the experiments during the light phase. To avoid the potential impact of hormonal variations,21 we did not use female animals. The study was reviewed and approved in accordance with the European Community directive 2010/63/UE and Spanish law regulating animal research (RD 1201/2005). All experimental protocols were reviewed and authorized by the Ethical Committee of the University of Cádiz for animal care and use.
Administration of Bromodeoxyuridine
Bromodeoxyuridine (BrdU) is incorporated into the newly synthesized DNA in actively dividing cells. Once incorporated, it remains in the nucleus of the cell and can be detected in fixed tissue by immunohistochemistry with specific antibodies. To measure the effect of chronic pain and/or stress on cell proliferation, as well as the survival and differentiation of postmitotic cells, we administered BrdU to rats according to the different protocols (Fig. 1A). In the proliferation protocol, animals received cumulative daily injections of BrdU over 5 days (100 mg/kg/d intraperitoneally [IP] in 0.9 % saline: Sigma-Aldrich Co., Madrid, Spain), starting 48 hours after surgery, and were killed 24 hours after the last injection. In the differentiation/survival protocol, BrdU was injected every 2 (50 mg/kg IP to give a total dose of 200 mg/kg) and 18 hours after the last BrdU injection, and the animals were exposed to the stress protocols over 4 weeks. During this survival period, labeled cells had extended axons,22 and they had begun to express mature neuronal markers.15,23
Neuropathic Pain Model: Chronic Constriction Injury
Chronic constriction injury (CCI) was induced as described previously.24,25 After we anaesthetized rats with an IP injection of sodium pentobarbital (50 mg/kg), the left sciatic nerve was exposed at the mid-thigh level, proximal to the sciatic trifurcation. Four chromic gut (4/0) ligatures (1.0–1.5 mm apart) were tied loosely around the nerve so as not to compromise vascular supply, and the overlying muscle and skin were then sutured with silk thread. Sham operations were performed in the same manner without ligating the nerve. The overall health of the animals was monitored regularly during the experimental period (weight, hair loss, and food and water intake) and animals displaying postsurgical complications or self-mutilation of at least 1 distal phalange on the operated paw were excluded from the study.
Behavioral Assessment of Pain
Mechanical allodynia and hyperalgesia were tested in rats before and at several time points after surgery (Fig. 1A). The animals were habituated to their environment for 15 minutes before behavioral testing, and each test was separated from the next by a 30-minute rest period. Mechanical allodynia was measured with an electronic version of the von Frey test (Dynamic Plantar Aesthesiometer, Ugo Basile, Gemonio, Varese, Italy), whereby a vertical force was applied to the left hind paw that increased from 0 to 50 g during a period of 20 seconds.26 Mechanical hyperalgesia was assessed by the paw-pinch test,27 which involved gradually applying increasing pressure via a graded motor-driven device (Ugo Basile) and establishing a 250-g cutoff to prevent tissue damage. In both tests, the pressure evoking a clear voluntary hind paw withdrawal response was recorded and taken as the threshold of mechanical nociception.
Once daily over 7 (proliferation experiment) or 28 days (differentiation experiment), rats were immobilized for 45 min in restraint bags (Rodent Restraint Cones; Harvard Apparatus, Holliston, MA). Unstressed animals remained in their boxes and were otherwise handled identically to the stressed animals.
Spontaneous locomotor activity was recorded individually for each animal 4 weeks after surgery. Rats were placed in the center of a Plexiglas, open-field square box (45 × 45 × 35 cm) and free exploratory behavior was monitored over 15 minutes by the use a camera connected to SMART video system (Spontaneous Motor Activity Recording and Tracking, Panlab S.L.U., Spain). Horizontal activity was analyzed over 5 minutes, and the total distance travelled was expressed in arbitrary units with respect to time.
Corticosterone was measured in plasma 8 days after surgery.28 Animals were anesthetized with 8% chloral hydrate (500 mg/kg, IP) and immediately after decapitation, and blood was collected in sodium di-hydrogen citrate (3.15 %) in a 0.1 M phosphate buffer as an anticoagulant. The blood samples were centrifuged at 1000g for 10 minutes at 4°C, and the plasma collected was stored at −80°C. Plasma corticosterone levels were assayed with a commercially available radioassay (Siemens Healthcare Diagnostics SL, Getafe, Spain), and a γ counter was used to measure the radioactivity incorporated (Wallac Wizard 1470, Perkin Elmer, Waltham, MA). The detection limit for this kit is 5.7 ng/mL, the intra-assay coefficient of variation was 4.3%, and the interassay coefficient of variation was 5.8%.
Tissue Treatment and Immunostaining
Rats were deeply anaesthetized with chloral hydrate (8%) and transcardially perfused with 4% (w/v) paraformaldehyde in 0.1 M phosphate buffer (pH 7.4). The rat’s brain was removed, postfixed for 2 hours, and cryoprotected by immersion in a 30% sucrose solution overnight. Coronal sections of the hippocampus (30-μm thick) were obtained on a cryostat and stored at −20°C in cryoprotectant solution (glycerol and phosphate-buffered saline [PBS; pH 7.4], 1:1 v/v). Free-floating cerebral sections were immunostained for ki67, BrdU, glial fibrillary acidic protein (GFAP), neuronal-specific nuclear protein (NeuN), or doublecortin (DCX), using the following primary antibodies: rabbit anti-ki67 (proliferation marker, 1:1000; Vector Laboratories Ltd., Peterborough, United Kingdom), rat anti-BrdU (1:100; Abcam, Cambridge, United Kingdom), rabbit anti-GFAP (glial marker, 1:5000; Dako Diagnosticos SA, Sant Just Desvern, Barcelona, Spain), goat anti-DCX (neuroblast marker, 1:100; Santa Cruz Biotechnology, Inc., Heidelberg, Germany), rabbit anti-Iba1 (microglia marker, 1:1000; Wako Pure Chemical Industries, Osaka, Japan), and mouse anti-NeuN (neuronal marker, 1:50; Chemicon, a Merck Millipore Company, Billerica, MA). To detect incorporation of BrdU, DNA was first denatured in free-floating cerebral sections by treatment with a solution containing 50% (v/v) formamide, 150 mM NaCl, and 15 mM sodium citrate at 65°C for 2 hours, followed by a 30-minute incubation in 2 N HCl at 37°C. For single labeling of BrdU or DCX, sections were first treated with 2% H2O2 and 60% (v/v) methanol in PBS for 30 minutes to block endogenous peroxidase activity. An antirat or antigoat biotinylated secondary antibody was used, and antibody binding was revealed using the avidin/biotinylated enzyme complex kit (Pierce, Thermo Scientific, Rockford, IL). The peroxidase reaction was visualized with diaminobenzidine (0.25 mg/mL) in a solution containing hydrogen peroxide (0.003 % v/v). For double or triple labeling, Alexa 647, 633, 488, or 565 conjugated secondary antibodies were used (Molecular Probes, Eugene, OR).
To prevent nonspecific antibody binding, sections were incubated for 30 minutes with a solution containing 2.5% (w/v) bovine serum albumin, 0.25% (w/v) sodium azide, and 0.1% (v/v) Triton X-100 in PBS. This solution was also used to dilute the primary and secondary antibodies. After staining, the sections were mounted on slides, dehydrated, coverslipped with DePeX, and analyzed by light microscopy. Alternatively, the sections were mounted with antifading Vectastain and analyzed by epifluorescent (BX60 epifluorescence microscope; Olympus, Barcelona, Spain) or confocal microscopy (Spectra confocal microscope, Leica, Barcelona, Spain). Optical images were captured with a digital camera (DP71: Olympus).
DG sections were obtained 1.2 and 3.8 caudal to bregma, according to the Rat Brain Atlas.29 Proliferating cells were identified by optical microscopy using a 40× objective (Optika Microscopes, Ponteranica, Italy), and unbiased stereologic methods were used to assess BrdU+ nuclei and ki67 immunostaining. In the DG, nuclei were counted in the hilus, subgranular zone, and the granule cell layer of every ninth section, and the data were expressed as the total number per DG. BrdU-labeled cells in the granule cell layer and subgranular zone of the DG were counted using a light microscope at 40× magnification by an experimenter blind to the treatment. Cells were considered part of the subgranular zone of the DG if they bordered or fell within the granular layer. The cell counts from each section were added and multiplied by 9 to obtain the total number of BrdU-labeled cells in the DG for each brain. The number of DCX+ cells was estimated by analyzing 8 sections per animal and 5 to 6 animals in each condition. The percentage of NeuN+ and GFAP+ or iba1+ nuclei was quantified by confocal microscopy as the number of labeled nuclei that also incorporated BrdU among a total of 50 to 60 BrdU+ nuclei from 3 animals per experimental group.30 To calculate the total number of nuclei with BrdU and NeuN, the percentage colocalization was multiplied by the number of BrdU+ nuclei in each animal.
The rats were distributed on 3 independent experimental sets: (i) nociceptive behavioral tests, (ii) corticosterone evaluation, and (iii) motor activity test. Tissues from animals used to explore spontaneous motor activity subsequently were used to perform inmunohistochemistry studies (single- or double-cell staining). The exact number of animals per group for each group is detailed in the Results and Tables 1 and 4. All data are presented as the mean ± SEM, and all the results were analyzed using SPSS 21 (SPSS, Chicago, IL) and STATISTICA 10.0 (StatSoft, Tulsa, OK) software. The data were tested for normality and homogeneity of variance with the Kolmogorov-Smirnov normality test and Levene test for equal variances, respectively. Data that satisfied these assumptions (P > 0.05) were subsequently analyzed by 1- or 2-way analysis of variance (ANOVA) and post hoc Bonferroni test for multiple comparisons (Bonferroni corrected). Otherwise, nonparametric methods (Kruskal-Wallis test) were used. The level of significance was set at a P value of <0.05. P value <0.01 was used when n ≤ 3. Detailed information regarding statistical analysis is shown in Tables 1–6.
Assessment of Pain, Corticosterones, and Locomotor Activity After Neuropathic Pain and Stress
Animals were separated into 2 experimental groups for the proliferation and differentiation/survival experiments (Fig. 1A). All CCI rats developed a typical pain phenotype, evident through their flinching and the paw guarding posture adopted. In the proliferation experiments, rats were tested for the presence of mechanical allodynia at several time points after surgery (von Frey test: Fig. 1B). Five days after the experimental induction of neuropathy, CCI, and CCI + stress, rats (n = 5 per group) exhibited pronounced mechanical allodynia of the operated hind paw (2-way ANOVA: FCCI (1,15) =91.2, P = 0.0001; Fstress (1,15) = 3.6; FCCI × stress (1,15) = 0.36: Fig. 1B; Tables 1–3). In this test, the threshold decreased by up to 5 to 10 g, and it remained at this level until the end of the experimental period.
In the differentiation/survival experiments, mechanical hyperalgesia was evaluated (paw pinch test: Fig. 1C), and neuropathic animals developed strong mechanical hyperalgesia after surgery that persisted for at least 4 weeks (2-way ANOVA: FCCI (1,15) = 49.9, P = 0.0001; Fstress (1,15) = 2.25; FCCI × Stress (1,15) = 0.05: Fig. 1C; Tables 1–3). Although CCI animals displayed a permanent limp, they showed no significant differences in the speed (data not shown) or total distance traveled in the test 4 weeks after the induction of chronic pain (1-way ANOVA: F(3,26) = 2.6, P > 0.05: Fig. 1D; Tables 4 and 5). Overall, the pain tests showed that CCI animals exhibited similar sensorial thresholds regardless of whether they were subjected to stress, suggesting that, contrary to earlier data, stress did not alter sensory pain.18
In addition, and as an index of stress, plasma corticosterone levels were measured in these rats 8 days after surgery. A significant increase in plasma corticosterone levels was detected in the stressed rats compared with the unstressed animals (1-way ANOVA: F(3,16) = 13.9, P < 0.0001: Fig. 1E; Tables 4–6), yet no such increase was produced in CCI rats, consistent with previous finding.31 Finally, no differences in body weight were observed among the distinct groups of rats (data not shown).
Neuropathic Pain Decreased Hippocampal Cell Proliferation, an Effect That Was Increased by Stress
To determine the effect of neuropathic pain and restraint stress on the number of dividing cells, cell proliferation in the DG was analyzed by immunohistochemistry when the animals were killed for Ki67 in animals perfused 1 week after surgery (proliferation experiment: Fig. 2A). This antibody detects cells in the process of dividing in fixed tissues, and although there were many areas with labeled cells, only the cells located in the subgranular zone were quantified (inset in Fig. 2A). The number of Ki67+ cells was significantly lower (Table 4) 1 week after surgery in animals who experienced neuropathy (57.3% ± 5.1%, n = 6) than in the sham group (113.3% ± 15.4%, n = 6) but not in stress group (86.1% ± 5.9%, n = 7). This reduction was exacerbated in the CCI + stress group (37.3% ± 7.3%, n = 7) and statistically significant (1-way ANOVA: F(3,22) = 13.5, P < 0.0001, Tables 5 and 6) compared with the sham (P < 0.0001), stress (P = 0.004), and CCI group (P = 0.048). When the dorsal and ventral hippocampus was quantified separately, there were no significant differences between these regions (data not shown).
Neuropathic Pain and Concomitant Restraint Stress Decrease Neuroblast Number in the DG
To verify that the alterations in proliferation affected cells of a neuronal lineage, we examined the DCX-expressing cells in the DG of the hippocampus in animals subjected to neuropathy and/or restraint stress for 4 weeks. DCX is an intermediate filament protein expressed by neuroblasts <1 month old.32 Therefore, this marker labeled most neuroblasts that proliferated and that began to differentiate during the exposure to pain and/or restraint stress. These cells were located in the subgranular zone, and they had abundant projections extending throughout the granular layer (Fig. 3A). When the data were quantified, both pain (62.0 ± 6.7 DCX+ cells/slice, P = 0.006, n = 6) and immobilization stress (60.9 ± 4.5 DCX+ cells/slice, P = 0.007, n = 5) reduced the number of neuroblasts in the hippocampal DG area compared with sham-operated rats (93.6 ± 4.5 DCX+ cells/slice, n = 6; 1-way ANOVA: F(3,18) = 13.7, P < 0.0001, Fig. 3B, Tables 4–6). Moreover, this effect was exacerbated when CCI and stress were combined (40.3 ± 5.1 DCX+ cells/slice, P < 0.02, n = 5). These results indicate that the antiproliferative effect of pain and stress affects neuronal lineages.
Neuropathic Pain Decreases the Survival of Postmitotic Neurons in the DG, an Effect Enhanced by Restraint Stress
To determine whether pain and restraint stress affected postmitotic cell survival, adult rats were injected with BrdU before surgery and killed 4 weeks later. BrdU was incorporated into cells located in the subgranular zone (survival experiment: Fig. 4A) and embedded in the granular cell layer (inset in Fig. 4A), and there were significant differences in the number of postmitotic cells between groups (1-way ANOVA: F(3,18) = 10.22, P < 0.0001, Fig. 4B, Tables 4–6). Indeed, the number of postmitotic cells decreased significantly in animals subjected to CCI (4458 ± 374, P = 0.01, n = 5) but not to restraint stress (5290 ± 1094, P = 0.11, n = 5) compared with sham animals (7696 ± 527, n = 6). Furthermore, in animals subjected to both CCI and immobilization stress, the decrease in the number of postmitotic cells per animal was even greater, as large as a approximately 65% reduction (2737 ± 86, P < 0.0001, n = 6: Fig. 4B).
To determine the phenotype of the cells that incorporated BrdU, parallel series of sections from the same animals were used for BrdU, NeuN, and GFAP triple labeling, or for Iba1 and BrdU double labeling. When the images were analyzed by confocal microscopy (Fig. 4, C and D), very few GFAP+/BrdU+ or Iba1+/BrdU+ cells were detected (<2%), with no significant differences between the groups (data not shown). In most of the nuclei that incorporated BrdU (~70%), NeuN was coexpressed (BrdU+/NeuN+ nuclei). The percentage of BrdU+ cells coexpressing NeuN was similar in all the animals (1-way ANOVA: F(3,8) = 2.2, P = 0.17: Fig. 4C; Tables 4–6), indicating that neither CCI nor immobilization stress altered postmitotic differentiation. The density of newborn cells expressing each combination of markers in the subgranular zone and granular layer (absolute number of BrdU+/NeuN+ cells) was determined by multiplying the percentages of BrdU+/NeuN+ cells by the overall density of BrdU+ cells in each region per animal (sham animals 4754 ± 286, n = 3: Fig. 4E). Animals subjected to CCI or restraint stress did not exhibit a decrease in the number of postmitotic cells expressing NeuN (Kruskal-Wallis test, H(3,12) = 8.12, P = 0.04; CCI, 2886 ± 189, P = 1, n = 3; stress group, 3053 ± 745, n = 3; P = 0.68), and this reduction was statistically significant only in CCI + stress rats, reaching 1889 ± 230 BrdU+ neurons (P = 0.009, n = 3, Table 6).
Although decreased hippocampal neurogenesis is observed in many models of stress, the effects of neuropathic pain on neurogenesis have not been studied extensively. Indeed, the potential influence of stress on neurogenesis when associated with neuropathic pain remains unclear, despite the importance of this factor in explaining some of the conditions that may develop in association with pain, such as anxiety, depression, and cognitive decline. In this study, we examined proliferation and survival of neuroblasts into dentate gyrus of rats exhibiting marked mechanical allodynia and hyperalgesia, which would demonstrate that neuropathic pain inhibits these phenomena in the hippocampus. Indeed, neuropathic pain diminishes the number of neuroblasts and the incorporation of new neurons into the granule cell layer. Moreover, this antineurogenic effect of chronic pain is increased by concomitant low-intensity stress.
We found a reduction in the proliferation rate in the DG, evident through Ki67 labeling, 1 week after neuropathy is induced. This reduced proliferation led to a decrease in the total number of neuroblasts (DCX + cells) after 4 weeks, in agreement with previous data showing that neuropathic pain lasting for 2 weeks8,17 or 35 days10 decreased the number of neuroblasts in the hippocampus of adult mice. Furthermore, it is noteworthy that the distribution of postmitotic cell phenotypes is not modified by chronic pain, because most cells differentiate into neurons and a small percentage of them into glial cells. Besides, our work shows for the first time that this decrease in the number of proliferating cells implies that fewer new neurons integrate into the granule cell layer of the hippocampus. This is evident through the number of BrdU+/NeuN+ cells found after 4 weeks of BrdU administration, indicating that CCI specifically decreases neurogenesis, but it does not seem to affect gliogenesis.
Stress is one of the most potent environmental factors known to suppress adult neurogenesis, as evident in different species and using several stress paradigms.33,34 With the protocol used in our experiments (45 minutes restraint/day), we tried to mimic the situation of patients having to deal with daily stressful situations unrelated to their chronic pain pathology. The restraint stress protocol used here reduced the number of neuroblasts, as well as cell proliferation or survival, while increasing the blood corticosterone levels. The association between chronic high levels of glucocorticoids and decreased neurogenesis is well established.35–39 Thus, the high levels of corticosterone detected may have contributed to the impaired maturation and proliferation of neurons in stressed animals, but not that in CCI animals that have been shown to have normal levels of this glucocorticoid.
This finding is consistent with previous studies,10,40–43 where no changes in plasma corticosterone were detected in animals suffering neuropathic pain. Thus, the effect of neuropathic pain on neurogenesis cannot be explained on the basis of a negative experience, obviously stressful, that is intrinsically associated with pain. That is, neuropathic pain per se seems to differ from stress, at least for the neuroendocrine response, even if it induces similar consequences at the hippocampal level. Conversely, it is widely known that stressful experiences44 and neuropathic pain condition45,46 lead to sleep disturbances, and it has been proposed that pattern and duration of sleep have an important effect on the rate neurogenesis.47,48 Therefore, it is likely that a sleep-related alteration may be an underlying, hippocampal dysfunction.
Our study also explores the hippocampal consequences of adding stress to a painful experience. This is a matter of particular interest, because many studies have shown that stressful experiences negatively influence pain perception, although the underlying biological mechanisms remain unclear. Our data show that stress increases the antiproliferative effect of pain and diminishes the survival of postmitotic neurons in the DG. Therefore, when stress and pain coexist, the inhibition of neurogenesis is exacerbated, suggesting a summation of the effects of both conditions. Furthermore, it is interesting to note that in these animals, corticosterone levels remain as high as in the stress group, which suggests that the combination of neuropathic pain and stress does not provoke a more severe alteration in hypothalamo-pituitary-adrenal function.
The continuous decrease in the number of new neurons incorporated into the granular cell layer of the DG might produce synaptic and functional changes in the hippocampus of animals suffering neuropathic pain, which may be more robust in those that also are subjected to stress. In the hippocampal circuit, each granule cell contacts approximately 10 to 15 CA3 pyramidal neurons,49 and they project their axons to the CA1. Therefore, the consequences of a reduction in the number of these cells may explain the inhibition of long-term potentiation at CA3 to CA1 synapses and the reduction of CA1 synaptic boutons in the hippocampus after peripheral nerve injury.4,6 Similar alterations in mossy fiber-CA3 plasticity have been described in the hippocampus of mice with neuropathy.8 Furthermore, the reduction in neurogenesis may also involve proinflammatory cytokines, such as tumor necrosis factor-α,4 or others found to be affected in the hippocampus postperipheral injury.50,51 In situ hybridization assay results demonstrated that either pain or stress (acute or chronic treatments) reduced the levels of both neurokinin-1 receptor and brain-derived neurotrophic factor mRNAs in the hippocampus, and brain-derived neurotrophic factor is a potent activator of neurogenesis.12 Conversely, the activation of N-methyl-D-aspartate receptors by glutamate is essential for both normal neurogenesis52,53 and the development of neuropathic pain states (central sensitization).54 All these changes could explain the deficits in the functionality of this structure that occurs during chronic pain. Indeed, a reduction in spatial memory is associated with neuropathic pain in rodent models 4 weeks after surgery.4,55,56 We propose that this cognitive impairment may be mediated, at least in part, by the decrease in the number of new neurons in the DG.
Besides memory and learning process, hippocampus malfunction has been implicated widely in affective disorders. In this sense, neuropathic pain induces anxiety and depression-related behaviors in a time-dependent manner. Indeed, neuropathic rats developed anxiety and depression-related behaviors 4 weeks after the induction of neuropathic pain in our laboratory7 These results were obtained in a comparable time frame and suggest that affective pathologies related with pain require long-term molecular and neural plasticity.57 Support for the neurogenic hypothesis of depression has come from studies showing that although depression states inhibit neurogenesis,58,59 many classes of antidepressants enhance such processes in the subgranular zone of the hippocampus.60–63 In addition, and despite a few recent exceptions,64,65 a reversal of depressive-like behavior by antidepressants is associated with hippocampal neurogenesis.62,66–69 Moreover, a proneurogenic effect of pregabalin was recently described in adult mice exposed to chronic stress.70
Taking into account all these results, we speculate that pain-related decreases in neurogenesis may promote, at least in part, the emergence of depressive symptoms in patients with chronic pain. It is important to note that the data presented do not permit a cause–effect relationship between the decrease in neurogenesis and the behavioral consequences to be established, which can be demonstrated only through further experiments. Finally, it has been shown that steroid hormones are important in regulating adult neurogenesis,21 and, as a consequence, adult neurogenesis in rats is a process differentially modulated in the 2 sexes. Therefore, further studies in female rats are necessary to know the influence of chronic pain and stress in neurogenesis.
In summary, our findings show that neuropathic pain decreases the proliferation and survival of newly formed neurons in the adult rat hippocampus. The most robust down-regulation of neurogenesis occurred in animals in which neuropathic pain co-exists with stress. We hypothesize that the cognitive deficits and mood disorders observed in patients with chronic pain would be at least partially due to the decrease in the number of new neurons incorporated into the hippocampus and that this deficit may be exacerbated in patients with stress. Additional experiments are needed to determine whether hippocampal neurogenesis affects the chronicity of pain per se or in pathologies associated with chronic pain, such as cognitive deficits and depression.
Name: Carmen Romero-Grimaldi, PhD.
Contribution: This author helped conduct the experiments (neurogenesis), design the experiments, and write the final manuscript.
Attestation: Carmen Romero-Grimaldi approved the final manuscript, attests to the integrity of the original data and the analysis reported in this manuscript, and is responsible for maintaining the study records.
Name: Esther Berrocoso, PhD.
Contribution: This author helped design the study and prepare the manuscript.
Attestation: Esther Berrocoso approved the final manuscript.
Name: Cristina Alba-Delgado, PhD.
Contribution: This author helped conduct the experiments (pain behavior).
Attestation: Cristina Alba-Delgado approved the final manuscript.
Name: Jose Luis M. Madrigal, PhD.
Contribution: This author helped conduct the experiments (corticosterone study).
Attestation: Jose Luis M. Madrigal approved the final manuscript.
Name: Beatriz G. Perez-Nievas, PhD.
Contribution: This author helped conduct the experiments (corticosterone study).
Attestation: Beatriz G. Perez-Nievas approved the final manuscript and attests to the integrity of the original data and the analysis reported in this manuscript.
Name: Juan Carlos Leza, MD, PhD.
Contribution: This author helped design the study and prepare the manuscript.
Attestation: Juan Carlos Leza approved the final manuscript and attests to the integrity of the original data and the analysis reported in this manuscript.
Name: Juan Antonio Mico, MD, PhD.
Contribution: This author helped design the study and prepare the manuscript.
Attestation: Juan Antonio Mico approved the final manuscript and attests to the integrity of the original data and the analysis reported in this manuscript.
This manuscript was handled by: Jianren Mao, MD, PhD.
We thank Mrs. Raquel Rey-Brea (Research Technician, CIBER of Mental Health (CIBERSAM), ISCIII, Madrid, Spain) and Mrs. Clara Muñoz-Mediavilla and Mr. Jose-Antonio Garcia-Partida (Research Technician at the Department of Neuroscience [Pharmacology and Psychiatry], University of Cádiz, Cádiz, Spain) for their excellent technical assistance.
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