Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2019 Jun 1.
Published in final edited form as: Exp Neurol. 2018 Mar 9;304:125–131. doi: 10.1016/j.expneurol.2018.03.005

DREADDed microglia in pain: implications for spinal inflammatory signaling in male rats

Peter M Grace 1,2,5,*, Xiaohui Wang 1,3,6,*, Keith A Strand 1, Michael V Baratta 1,*, Yingning Zhang 1, Erika L Galer 1, Hang Yin 3, Steven F Maier 1, Linda R Watkins 1
PMCID: PMC5916033  NIHMSID: NIHMS951471  PMID: 29530713

Abstract

The absence of selective pharmacological tools is a major barrier to the in vivo study of microglia. To address this issue, we developed a Gq- and Gi-coupled Designer Receptor Exclusively Activated by a Designer Drug (DREADD) to enable selective stimulation or inhibition of microglia, respectively. DREADDs under a CD68 (microglia/macrophage) promoter were intrathecally transfected via an AAV9 vector. Naïve male rats intrathecally transfected with Gq (stimulatory) DREADDs exhibited significant allodynia following intrathecal administration of the DREADD-selective ligand clozapine-N-oxide (CNO), which was abolished by intrathecal interleukin-1 receptor antagonist. Chronic constriction injury-induced allodynia was attenuated by intrathecal CNO in male rats intrathecally transfected with Gi (inhibitory) DREADDs. To explore mechanisms, BV2 cells were stably transfected with Gq or Gi DREADDs in vitro. CNO treatment induced pro-inflammatory mediator production per se from cells expressing Gq-DREADDs, and inhibited lipopolysaccharide- and CCL2-induced inflammatory signaling from cells expressing Gi-DREADDs. These studies are the first to manipulate microglia function using DREADDs, which allow the role of glia in pain to be conclusively demonstrated, unconfounded by neuronal off-target effects that exist for all other drugs that also inhibit glia. Hence, these studies are the first to conclusively demonstrate that in vivo stimulation of resident spinal microglia in intact spinal cord is a) sufficient for allodynia, and b) necessary for allodynia induced by peripheral nerve injury. DREADDs are a unique tool to selectively explore the physiological and pathological role of microglia in vivo.

Keywords: Minocycline, neuropathic pain, chemogenetics, intrathecal, allodynia, AAV, gene therapy, CNO

Introduction

Microglia, the resident macrophages of the central nervous system (CNS), have a key role in maintaining homeostasis and in driving many CNS pathologies, including persistent pain (Grace et al., 2014; Ji et al., 2016; Old et al., 2015; Salter and Stevens, 2017). Precise delineation of their function in vivo has been hindered by a lack of tools to selectively activate or inhibit microglia. There are no strategies in routine use to selectively activate microglia without activating other cell types as well. In addition, several approaches have been used to inhibit microglia, but all are flawed with respect to specificity. For example, pharmacological agents such as minocycline, which do inhibit microglia (Yrjänheikki et al., 1998), also have inhibitory effects on other cells in the CNS like neurons, astrocytes and T cells (e.g., (González et al., 2007; Song et al., 2016; Szeto et al., 2011)). Selective depletion methods have been described, such as CSF-1R antagonism (Elmore et al., 2014), or cre-driven expression of the diphtheria toxin receptor in microglia (Parkhurst et al., 2013). However, these methods widely perturb the CNS, as cytokine production and/or astrogliosis is reported. These off-target effects may confound interpretations regarding the role of microglia in physiological and pathological processes. We have recently reported that hM4Di Designer Receptors Exclusively Activated by Designer Drugs (DREADDs) can be used to inhibit microglia in vitro and in vivo (Grace et al., 2016a). Once expressed under a cell-specific promoter, DREADDs can be remotely activated with clozapine-N-oxide (CNO) (Armbruster et al., 2007). Gi-linked signaling was predicted to attenuate microglial reactivity because activation of the M4 muscarinic receptor (the Gi DREADD progenitor (Armbruster et al., 2007)) inhibits Ca2+ influx in parasympathetic neurons (Cuevas and Adams, 1997), a process associated with decreased proinflammatory cytokine production in microglia (Hayashi et al., 2011; Hoffmann et al., 2003). When spinal microglia were transfected with hM4Di DREADDs under a CD68 promoter in vivo, treatment with CNO prevented and reversed morphine-induced persistent sensitization in male rats (Grace et al., 2016a). CNO also attenuated inflammatory gene expression in hM4Di+ BV-2 cells (a microglia cell line) that were stimulated with the danger signal HMGB1 (Grace et al., 2016a; Lacagnina et al., 2017). Inflammatory mediators, such as nitric oxide (NO) and interleukin-1β (IL-1β) promote neuronal hyperexcitability in the spinal dorsal horn (Grace et al., 2014, 2016b). However, it is not known whether these results apply to neuropathic pain per se, and extend to other BV-2 cell stimuli in vitro.

hM3Dq DREADDs have also been developed that mobilize intracellular calcium through Gq proteins (Armbruster et al., 2007). As noted above, increased intracellular Ca2+ is a substrate for microglial activation and cytokine release (Hoffmann et al., 2003). Whether hM3Dq DREADDs can activate microglia is unknown.

The first aim of this study was to determine whether activation or inhibition of microglia via DREADDs could influence nociceptive hypersensitivity. The second aim was to validate behavioral results in microglia cell cultures by determining whether DREADD activation or inhibition would influence cytokine transcription and translation. Our data indicate that DREADDs can be used to modulate microglial activity in vitro and in vivo.

Materials and methods

Subjects

Pathogen-free adult male Sprague Dawley rats (n = 6 rats/group for each experiment; 10–12 wks old on arrival; Envigo Labs, Indianapolis, IN) were used in all experiments. Rats were housed in temperature-controlled (23 ± 3°C) and light-controlled (12 h light:dark cycle; lights on at 07:00 h) rooms with standard rodent chow and water available ad libitum. All procedures were approved by the Institutional Animal Care and Use Committee of the University of Colorado Boulder.

DREADDs and drugs

pAAV-CD68-hM4Di-mCherry, pAAV-CD68-hM3Dq-mCherry, pAAV-CD68-EGFP (control), pLenti6.3-CMV hM4Di, and pLenti6.3-CMV hM3Dq plasmids were generated as previously described (Grace et al., 2016a) (gifted by D.J. Urban and B.L. Roth). pAAV plasmids were packaged in serotype 9 by the UNC Vector Core Facility. Clozapine-N-oxide (CNO; Enzo Life Sciences, Farmingdale, NY) was obtained commercially, while IL-1ra was gifted by Amgen (Thousand Oaks, CA). Vehicles were DMSO (1.5% vol/vol) for CNO and sterile saline for IL-1ra.

Chronic constriction injury (CCI) surgery

Neuropathic pain was induced using the CCI model of sciatic nerve injury (Bennett and Xie, 1988). Surgery was performed at the mid-thigh level of the left hindleg as previously described (Grace et al., 2010). In brief, animals were anesthetized with isoflurane. The shaved skin was treated with Nolvasan and the surgery was performed aseptically. Four sterile chromic gut sutures (cuticular 4-0 WebGut, Patterson Veterinary, Devens, MA) were loosely tied around the gently isolated sciatic nerve. Animals were monitored post-operatively until fully ambulatory, prior to return to their home cage.

Mechanical allodynia

All testing was conducted blind with respect to group assignment. Rats received at least three 60-minute habituations to the test environment prior to behavioral testing. The von Frey test (Chaplan et al., 1994) was performed as previously described (Grace et al., 2010). A logarithmic series of 10 calibrated Semmes-Weinstein monofilaments (von Frey hairs; Stoelting, Wood Dale, IL) were applied randomly to the left versus right hind paws to define the threshold stimulus intensity required to elicit a paw withdrawal response. Log stiffness of the hairs ranged from manufacturer-designated 3.61 (0.40 g) to 5.18 (15.14 g) filaments. The behavioral responses were used to calculate absolute threshold (the 50% probability of response) by fitting a Gaussian integral psychometric function using a maximum-likelihood fitting method (Harvey, 1986; Treutwein and Strasburger, 1999).

Acute and chronic catheter implantation

The method of construction and implantation of the acute and indwelling intrathecal catheters was based on that described previously (Milligan et al., 1999). In brief, intrathecal operations were conducted under isoflurane anesthesia by threading sterile polyethylene-10 tubing (PE-10 Intramedic Tubing; Becton Dickinson Primary Care Diagnostics, Sparks, MD, USA) guided by an 18-gauge needle between the L5 and L6 vertebrae. The catheter was inserted such that the proximal catheter tip lay over the lumbosacral enlargement. For acute intrathecal drug delivery, catheters were removed immediately following drug delivery. For indwelling catheters, the needle was removed and the catheter was sutured to the superficial musculature of the lower back. The catheters were 17 cm in length, and were attached to a pre-loaded osmotic minipump where appropriate (Alzet, 2001, Cupertino, CA).

Intrathecal and intraperitoneal drug administration

For acute intrathecal drug delivery, the catheters were pre-loaded with drugs at the distal end in a total volume and delivered over 20–30 s once the catheter was in position. Acute intrathecal doses were as follows, DREADDs: 1.2 × 1013 vg in 8 μl (injected 4 weeks prior to CNO treatment); CNO 60 μg in 10 μl; IL-1ra: 100 μg in 10 μl. For repeated CNO treatment, 60 μg in 10 μl was delivered daily for 3 days. Intraperitoneal CNO was administered at 1 mg/kg. We have previously validated microglial expression of DREADDs when transfected via an AAV9 vector (Grace et al., 2016a).

Immunohistochemistry

After 3 days of CNO treatment, rats were deeply anesthetized with sodium pentobarbital and transcardially perfused with saline, followed by 4% paraformaldehyde/0.1 M phosphate buffer. Spinal cords were dissected and postfixed for 24 h in 4% paraformaldehyde/0.1 M phosphate buffer, and cryoprotected in 30% sucrose with 0.1% azide at 4°C until slicing. Lumbar spinal cords were freeze-mounted in OCT, and cut at 20 μm frozen sections. Sections were permeabilized with 0.3% hydrogen peroxide, blocked for 1 h with 10% NGS, 0.3% Triton-X in PBS, and then incubated overnight at 4°C for 24 h in 2 % normal goat serum together with mouse anti-rat CD11b (1:1000; Bio-Rad, Hercules, CA). Slides were then washed, incubated in the secondary antibody at 1:200 (Goat anti mouse biotin, Jackson Immuno Research, West Grove, PA) for 2 h. Sections were once again washed, incubated in ABC solution (Vector Laboratories, Burlingame, CA) for 2 h, washed, and incubated in inactive DAB (Sigma) for 10 min. DAB was then activated with B–D glucose (10mg/ml) and slides were incubated for 8 min, washed, and dried overnight. Slides were dehydrated in increasing concentrations of EtOH (50, 70, 95 and 100%), cleared in Citrisolv, dried and covered with DPX mountant (Sigma).

Images were acquired using an Olympus BX61 microscope (Olympus, Center Valley, PA) with Suite Cell Sens Dimension software (Olympus). All images were taken using the same exposure and settings, and captured at 20x magnification. Images were converted to 32 bit, corrected for threshold, and densitometry was conducted using NIH Image J software, while blinded to treatment conditions. Four images per animal were taken with 3 selections within each image analyzed, resulting in 12 areas of analysis per animal. Data are expressed as total area positive for staining within the selected area.

BV-2 transfection, cell culture, and treatments

Mouse microglia BV-2 cells were grown in DMEM medium supplemented with 10% FBS. Cells were seeded at a density of 1 × 104 cells/ml in 35 mm dish. When cells reached ~60% confluence, the medium was aspirated and 0.5 ml of pLenti6.3-CMV hM3Dq or pLenti6.3-CMV hM4Di lenti-virus medium (multiplicity of infection=104) and 8 μg/ml polybrene were added for transfection. After 24 h transfection, the virus was removed and the cells were cultured in DMEM medium supplemented with 10% FBS for additional 24 h. The pLenti6.3-CMV hM3Dq or pLenti6.3-CMV hM4Di positive cells were then selected by DMEM medium supplemented with 10% FBS and 10 μg/ml puromycin. Positive colonies were picked up and were further cultured. The pLenti6.3-CMV hM3Dq and pLenti6.3-CMV hM4Di positive cells were further confirmed by V5 tag western blotting.

pLenti6.3-CMV hM3Dq or pLenti6.3-CMV hM4Di positive cells were seeded into 96 well plates at a density of 5 × 104 cells/ml in DMEM medium supplemented with 10% FBS. After 24 h, all cells were washed. pLenti6.3-CMV hM3Dq cells were incubated with a concentration range of CNO (0, 50, 100 μM) in serum-free DMEM medium for 24 h. The CNO concentrations used did not adversely affect cell viability (Fig. S1). pLenti6.3-CMV hM4Di cells were incubated with a concentration range of LPS (0, 20, 200, 2000 μg/ml) or CCL2 (0, 50, 100 μg/ml), together with 0 or 50 μM of CNO in serum-free DMEM medium for 4 or 24 h. Non-transfected were treated with a concentration range of CNO (0–400 μM), together with a LPS (0 or 200 ng/ml) in serum-free DMEM medium for 24 h. Cells were collected by centrifugation for RT-PCR, whereas supernatants were collected for NO and cytokine analysis.

RT-PCR

Total RNA was isolated using a standard method of phenol:chloroform extraction (Chomczynski and Sacchi, 1987). cDNA amplification was performed using Quantitect SYBR Green PCR kit (Qiagen) in iCycler iQ 96-well PCR plates (Bio-Rad) on a MyiQ single Color Real-Time PCR Detection System (Bio-Rad). Primer sequences (GenBank, National Center for Biotechnology Information; www.ncbi.nlm.nih.gov) are as follows, Gapdh: F: GGAGAAACCTGCCAAGTATG, R: GTCATTGAGAGCAATGCCAG; Nos2: F: GGAGTGACGGCAAACATGACT, R: TCGATGCACAACTGGGTGAAC; Il1b: F: TGCTGTCGGACCCATATGAG, R: ATCCACACTCTCCAGCTGCA. Each sample was measured in duplicate by using the MyiQ single Color Real-Time PCR Detection System (Bio-Rad). Threshold for detection of PCR product was set in the log-linear phase of amplification and the threshold cycle (CT) was determined for each reaction. The level of the target mRNA was quantified relative to the housekeeping gene (GAPDH) using the ΔΔCT method (Livak and Schmittgen, 2001). GAPDH was not significantly different between treatments.

NO assay

The NO (nitrite) concentration in the culture supernatant was determined by the 2,3-diaminonaphthalene-based fluorescent method (Misko et al., 1993). The plate was read on a Beckman Coulter DTX880 reader (λex 365 nm, λem 430 nm).

ELISA

A total of 50 μl of cell culture supernatants were used for the assays. Cytokine levels were determined using commercially available rat TNF, IL-1β, and IL-6 ELISA kits (R&D Systems). The assays were performed according to the manufacturer’s instructions. Bradford protein assays were also performed to determine total protein concentrations in sonication samples. The analyte concentrations are presented as nanograms per mg of total protein.

Cell viability assay

Crystal violet staining was used to determine cell viability as described previously (Selfridge et al., 2015; Wang et al., 2013). Twenty-four hours after treatment, cells were fixed by 3.7% paraformaldehyde (PFA) for 5 min and then stained by 0.05% crystal violet for 30 min. The plates were subsequently washed 2 times with tap water and dried for 30 min at room temperature. 200 μL of ethanol was added to each well, and the plates were shaken for 15 min at room temperature to dissolve the dye. Absorbance at 540 nm was measured with a Beckman Coulter DTX880 reader (Fullerton, CA, USA)

Statistics

Data from the von Frey test were analyzed as the interpolated 50% thresholds (absolute threshold) in log10 of stimulus intensity (monofilament stiffness in milligrams × 10). Baseline values were compared between groups using a one-way ANOVA. Differences between treatment groups were determined using repeated measures 2-way ANOVA, followed by Tukey or Sidak post hoc tests where appropriate. Differences in cytokine levels were determined using 1-way ANOVA or 2-way ANOVA, followed by Dunnett’s or Sidak post hoc tests, where appropriate. Differences in gene expression were determined using 2-way ANOVA and Sidak post hoc tests, while differences in densitometry were determined using 1-way ANOVA and Tukey post hoc tests. Statistical comparisons are indicated on the figures for clarity and ± standard error of the mean. Statistical significance was set at P < 0.05.

Results

hM4Di inhibition of microglia reverses neuropathic pain

To test whether DREADD inhibition of spinal microglia could reverse established neuropathic pain, rats were transfected with intrathecal CD68-hM4Di or the control vector CD68-eGFP. We have previously validated spinal microglial expression of DREADDs when transfected via an AAV9 vector (Grace et al., 2016a). Two weeks after CCI surgery, a single intrathecal dose of CNO selectively reversed allodynia in those rats expressing hM4Di versus control (Fig. 1a; time x treatment: F4,36 = 34.36, P < 0.001; time: F4,36 = 41.05, P < 0.001; treatment: F1,9 = 61.35, P < 0.001). Allodynia was rapidly reversed within 2 h (P < 0.001), remained reversed through 6 h (P < 0.001), and had returned by 24 h. Intraperitoneal CNO also reversed allodynia over a similar timecourse (data not shown).

Figure 1. Spinal microglia inhibition via CD68-hM4Di attenuates CCI-allodynia and CD11b upregulation in vivo.

Figure 1

Open in a new tab

(a) Naïve rats were transfected with intrathecal hM4Di or control DREADDs. Two weeks later, CCI was performed on all rats. After a further two weeks, all rats received a single intrathecal dose of CNO (60 μg). Von Frey thresholds were determined prior to CCI (baseline; BL), prior to CNO (0 h; 14 days post CCI), and across a 24 h timecourse after CNO injection. (b) CD11b density was quantified in the ipsilateral dorsal horn of lumbar spinal cords from naïve rats, nerve-injured rats expressing control DREADD and treated with CNO for 3 days (60 μg/day), and nerve-injured rats expressing hM4Di DREADD and treated with CNO for 3 days. Representative images of ipsilateral dorsal horn of lumbar spinal cord are shown for (c) naïve rats (d) nerve-injured rats expressing control DREADD and treated with CNO for 3 days, and (e) nerve-injured rats expressing hM4Di DREADD and treated with CNO for 3 days. Data are presented as mean ± SEM; n = 6/group; *P < 0.05, ***P < 0.001.

We quantified CD11b density in the lumbar ipsilateral dorsal horns (Fig. 1b–e; F2,9 = 33.45, P < 0.001). As expected, CD11b was upregulated by CCI in rats expressing the control construct and treated with CNO for 3 days, relative to naïve (P < 0.001). In comparison, 3 days of CNO treatment downregulated CD11b in rats with CCI and expressing the hM4Di DREADD (P < 0.05).

hM4Di inhibition of BV-2 microglia attenuates pro-inflammatory mediator production

The final experiments aimed to test whether inhibition of microglia could attenuate inflammation. BV-2 cells were transfected with hM4Di DREADDs and exposed to a concentration range of LPS (Fig. 2a–d). LPS activates TLR4, which has a wide-ranging role in pain (Lacagnina et al., 2017). CNO treatment attenuated expression of Nos2 mRNA (LPS treatment x CNO treatment: F3,32 = 9.00, P < 0.001; LPS treatment: F3,32 = 141.3, P < 0.001; CNO treatment: F1,32 = 22.67, P < 0.001) and NO (LPS treatment x CNO treatment: F3,32 = 65.28, P < 0.001; LPS treatment: F3,32 = 435, P < 0.001; CNO treatment: F1,32 = 230.5, P < 0.001), as well as Il1b mRNA (LPS treatment x CNO treatment: F3,32 = 13.4, P < 0.001; LPS treatment: F3,32 = 64.42, P < 0.001; CNO treatment: F1,32 = 39.09, P < 0.001) and IL-1β (LPS treatment x CNO treatment: F3,32 = 15.47, P < 0.001; LPS treatment: F3,32 = 120.7, P < 0.001; CNO treatment: F1,32 = 54.76, P < 0.001).

Figure 2. BV-2 cell inhibition via hM4Di attenuates LPS- and CCL2-induced inflammation.

Figure 2

Open in a new tab

BV-2 cells were transfected with hM4Di. (a–d) Cells were treated with a concentration range of LPS, together with 0 or 50 μM of CNO. After 4 h, (a) Nos2 or (c) Il1b mRNA was quantified from cell lysates. After 24 h, (b) NO or (d) IL-1β levels were quantified in supernatants. (e–h) Cells were treated with a concentration range of CCL2, together with 0 or 50 μM of CNO. After 4 h, (a) Nos2 or (c) Il1b mRNA was quantified from cell lysates. After 24 h, (b) NO or (d) IL-1β levels were quantified in supernatants. Data are presented as mean ± SEM; n = 5/group; ***P < 0.001.

To test whether hM4Di DREADDs could attenuate activation from a stimulus that was relevant to neuropathic pain, cells were treated with a concentration range of CCL2 (Fig. 2e–h). CCL2 is a putative neuron-to-microglia signal in neuropathic pain, as microglia express CCR2 (Abbadie et al., 2003; Bose et al., 2016; Feng et al., 2017; Tian et al., 2017; Xu et al., 2017). CNO treatment attenuated expression of Nos2 mRNA (CCL2 treatment x CNO treatment: F2,24 = 56.12, P < 0.001; CCL2 treatment: F2,24 = 71.6, P < 0.001; CNO treatment: F1,24 = 79.64, P < 0.001) and NO (CCL2 treatment x CNO treatment: F2,24 = 73.53, P < 0.001; CCL2 treatment: F2,24 = 113.7, P < 0.001; CNO treatment: F1,24 = 75.02, P < 0.001), as well as Il1b mRNA (CCL2 treatment x CNO treatment: F2,23 = 47.51, P < 0.001; CCL2 treatment: F2,24 = 47.04, P < 0.001; CNO treatment: F1,23 = 49.44, P < 0.001) and IL-1β (CCL2 treatment x CNO treatment: F2,24 = 10.02, P < 0.001; CCL2 treatment: F2,24 = 19.57, P < 0.001; CNO treatment: F1,24 = 6.954, P < 0.05). CNO had no independent effect on LPS-induced NO release in control BV-2 cells at the concentrations used (Fig. S1).

hM3Dq stimulation of microglia induces IL-1-dependent allodynia

This experiment aimed to test whether activation of spinal microglia with DREADDs is sufficient to induce hindpaw allodynia. Naïve rats were transfected with intrathecal CD68-hM3Dq or the control construct. A single intrathecal dose of CNO selectively induced allodynia in those rats expressing hM3Dq versus control (Fig. 3; time x treatment: F4,40 = 3.50, P < 0.05; time: F4,40 = 7.76, P < 0.001; treatment: F1,10 = 10.45, P < 0.01). Allodynia slowly developed over 4 h (P < 0.05), peaking at 6 h (P < 0.001), and had resolved by 24 h. Intraperitoneal CNO also induced allodynia over a similar timecourse (data not shown).

Figure 3. Spinal microglia stimulation via CD68-hM3Dq induces IL-1-dependent allodynia in vivo.

Figure 3

Open in a new tab

Naïve rats were transfected with intrathecal hM3Dq or control DREADDs. Four weeks later, all rats received a single intrathecal dose of CNO (60 μg). Intrathecal IL-1ra (100 μg) was administered 2 h after CNO. Von Frey thresholds were determined prior to (baseline; BL), and across a 24 h timecourse after CNO injection. Data are presented as mean ± SEM; n = 6/group; *P < 0.05, ***P < 0.001.

To test whether CD68-hM3Dq-mediated allodynia was dependent on IL-1, a prototypical pronociceptive cytokine, IL-1ra was administered 2 h after CNO. The drug administrations were timed to overlap the half-lives of CNO (~6 h) with IL-1ra (~4 h). IL-1ra administration completely prevented the induction of allodynia by hM3Dq/CNO (Fig. 3; time x treatment: F4,40 = 4.45, P < 0.01; time: F4,40 = 11.58, P < 0.001; treatment: F1,10 = 12.19, P < 0.01).

hM3Dq stimulation induces pro-inflammatory mediators in BV-2 cells

The next experiment aimed to test whether activation of microglia with DREADDs induces production of proinflammatory mediators that have a known role in pain. BV-2 cells were transfected with hM3Dq DREADDs and exposed to a concentration range of CNO. The CNO treatment dose-dependently increased release of NO (Fig. 4a; F2,12 = 100.8, P < 0.001), TNF (Fig. 4b; F2,12 = 13.21, P < 0.001), IL-1β (Fig. 4c; F2,11 = 23.46, P < 0.001) and IL-6 (Fig. 4d; F2,11 = 19.32, P < 0.001).

Figure 4. hM3Dq stimulation of BV-2 cells induces pro-inflammatory mediators in vitro.

Figure 4

Open in a new tab

BV-2 cells were transfected with hM3Dq, and treated with a concentration range of CNO for 24 h. Supernatants were collected and analyzed for (a) NO, (b) TNF, (c) IL-1β, (d) IL-6. Data are presented as mean ± SEM; n = 5/group; *P < 0.05, ***P < 0.001.

Discussion

We found that inhibition of spinal microglia via hM4Di DREADDs attenuated established allodynia and CD11b expression in the dorsal horn of the lumbar spinal cord after peripheral nerve injury. Furthermore, these DREADDs attenuated inflammatory signaling induced by LPS and CCL2 in vitro, complementing similar results with HMGB1 (Grace et al., 2016a). In contrast, activation of spinal microglia via hM3Dq DREADDs induced allodynia in naïve rats, which could be prevented by inhibiting IL-1 signaling. Activation of hM3Dq DREADDs also induced release of inflammatory mediators in vitro. The output of cytokines via hM3Dq is consistent with microglial activation (Grace et al., 2014; Ji et al., 2016). Thus, DREADDs may be a useful strategy to selectively activate microglia in vivo.

DREADDs were targeted to microglia using a CD68 promoter. CD68 was selected because of its short promoter sequence, which is optimal for vectors with limited packaging capacity such as AAV. CD68 is not uniquely expressed by microglia, but also by macrophages. Although recently challenged (Gu et al., 2016; Guan et al., 2016), monocytes may infiltrate the spinal dorsal horn after peripheral nerve injury (Echeverry et al., 2011; Sweitzer et al., 2002; Zhang et al., 2007). Therefore, we injected the DREADDs prior to CCI to exclude any possible contribution from leukocytes. However, this design does not restrict microglial proliferation, which was recently shown to contribute to nociceptive hypersensitivity after peripheral nerve injury (Gu et al., 2016). Because CD68 also has a specialized role in phagocytosis (Zotova et al., 2013), it is further possible that only a subset of microglia express DREADDs after transfection. While this strategy is still sufficient to induce behavioral changes, approaches that use more generic promoters, such as Iba1, may lead to wider expression. The recent development of cre-inducible DREADDs (Zhu et al., 2016) may also allow for microglia-selective targeting.

CNO was used to activate the DREADDs. It was recently reported that the metabolite clozapine is responsible for the activity of CNO in vivo (Gomez et al., 2017). Clozapine is anti-nociceptive, but at higher doses than those used here (Schreiber et al., 1999). Furthermore, our in vivo experimental design excludes an independent anti- or pro-nociceptive effect of CNO, as the control group received pAAV-CD68-EGFP and CNO treatment. Nevertheless, newer approaches have replaced CNO with other drugs, such as compound 21 and KORD, that have minimal off-target activity (Roth, 2016).

Finally, these data further support a fundamental role for microglia in pain in male rats. We show for the first time that selectively activating spinal microglia is sufficient to induce hindpaw allodynia. Furthermore, activation of microglia with DREADDs results in inflammation in vitro and in vivo, which is consistent with their response to microorganisms or to tissue damage (Salter and Stevens, 2017). Future studies may confirm these results in primary microglia cultures. While we show that in situ activation of microglia is sufficient to induce pain, others have adoptively transferred ex vivo activated microglia to validate a specific activation signal (Tsuda et al., 2003). We also show that microglia are necessary for neuropathic pain, as DREADD inhibition reverses allodynia in vivo, and attenuates inflammatory signaling to chemical challenge in vitro. The extent to which DREADD inhibition attenuates microglia proliferation and activated morphology may be investigated in future studies. The temporal control of DREADD activation through CNO is another advantage over other strategies like adoptive transfer. As recent studies have concluded that microglia are not participants in neuropathic pain in females (Sorge et al., 2015), DREADDs may be a useful tool to confirm these findings in the future.

Together with our prior study (Grace et al., 2016a), we report that hM3Dq and hM4Di DREADDs can be used to activate and inhibit microglia, respectively. Future studies may investigate the intracellular signaling pathways that are engaged by Gi and Gq DREADDs. Due to the selectivity of DREADDs, this tool may provide an advantage over other pharmacological and depletion methods. Therefore, DREADDs are a unique tool to selectively explore the physiological and pathological role of microglia in vivo.

Supplementary Material

supplement. Figure S1. Effect of CNO on control BV-2 cells.

BV-2 cells were treated with a concentration range of CNO and a fixed concentration of LPS. CNO suppressed LPS-induced NO levels at higher concentrations, and did not affect cell viability.

Highlights.

  • DREADDs under a CD68 promoter were intrathecally transfected via an AAV9 vector

  • Activation of microglia via Gq DREADDs induced allodynia

  • Inhibition of microglia via Gi DREADDs reversed neuropathic pain

  • Activation of BV-2 cells via Gq DREADDs induced proinflammatory cytokines

  • Inhibition of BV-2 cells via Gi DREADDs attenuated LPS- and CCL2-induced cytokines.

Acknowledgments

The authors gratefully acknowledge Dr. Daniel Urban and Dr. Bryan Roth who gifted the DREADDs. This work was supported by the American Pain Society Future Leaders in Pain Research Grants Program (P.M.G.); National Health and Medical Research Council CJ Martin Fellowship ID 1054091 (P.M.G.); American Australian Association Sir Keith Murdoch Fellowship (P.M.G.); National Natural Science Foundation of China Grants 21750110432 (P.M.G.), 21602216, and 21543013 (X.W.); National Key Research and Development Program of China grant 2016YFC0800907 (X.W.); Natural Science Foundation of Jilin Province grants 20160101211JC, 20160520045JH (X.W.); and, NIH Grants R01GM101279 (H.Y.), DE021966, and DA023132 (L.R.W.).

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

References

  1. Abbadie C, Lindia JA, Cumiskey AM, Peterson LB, Mudgett JS, Bayne EK, DeMartino JA, MacIntyre DE, Forrest MJ. Impaired neuropathic pain responses in mice lacking the chemokine receptor CCR2. Proc Natl Acad Sci U S A. 2003;100:7947–7952. doi: 10.1073/pnas.1331358100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Armbruster BN, Li X, Pausch MH, Herlitze S, Roth BL. Evolving the lock to fit the key to create a family of G protein-coupled receptors potently activated by an inert ligand. Proc Natl Acad Sci U S A. 2007;104:5163–5168. doi: 10.1073/pnas.0700293104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bennett GJ, Xie YK. A peripheral mononeuropathy in rat that produces disorders of pain sensation like those seen in man. Pain. 1988;33:87–107. doi: 10.1016/0304-3959(88)90209-6. [DOI] [PubMed] [Google Scholar]
  4. Bose S, Kim S, Oh Y, Moniruzzaman M, Lee G, Cho J. Effect of CCL2 on BV2 microglial cell migration: Involvement of probable signaling pathways. Cytokine. 2016;81:39–49. doi: 10.1016/j.cyto.2016.02.001. [DOI] [PubMed] [Google Scholar]
  5. Chaplan SR, Bach FW, Pogrel JW, Chung JM, Yaksh TL. Quantitative assessment of tactile allodynia in the rat paw. J Neurosci Methods. 1994;53:55–63. doi: 10.1016/0165-0270(94)90144-9. [DOI] [PubMed] [Google Scholar]
  6. Chomczynski P, Sacchi N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem. 1987;162:156–159. doi: 10.1006/abio.1987.9999. [DOI] [PubMed] [Google Scholar]
  7. Cuevas J, Adams DJ. M4 muscarinic receptor activation modulates calcium channel currents in rat intracardiac neurons. J Neurophysiol. 1997;78:1903–1912. doi: 10.1152/jn.1997.78.4.1903. [DOI] [PubMed] [Google Scholar]
  8. Echeverry S, Shi XQ, Rivest S, Zhang J. Peripheral nerve injury alters blood-spinal cord barrier functional and molecular integrity through a selective inflammatory pathway. J Neurosci Off J Soc Neurosci. 2011;31:10819–10828. doi: 10.1523/JNEUROSCI.1642-11.2011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Elmore MRP, Najafi AR, Koike MA, Dagher NN, Spangenberg EE, Rice RA, Kitazawa M, Matusow B, Nguyen H, West BL, et al. Colony-stimulating factor 1 receptor signaling is necessary for microglia viability, unmasking a microglia progenitor cell in the adult brain. Neuron. 2014;82:380–397. doi: 10.1016/j.neuron.2014.02.040. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Feng C, Wang X, Liu T, Zhang M, Xu G, Ni Y. Expression of CCL2 and its receptor in activation and migration of microglia and monocytes induced by photoreceptor apoptosis. Mol Vis. 2017;23:765–777. [PMC free article] [PubMed] [Google Scholar]
  11. Gomez JL, Bonaventura J, Lesniak W, Mathews WB, Sysa-Shah P, Rodriguez LA, Ellis RJ, Richie CT, Harvey BK, Dannals RF, et al. Chemogenetics revealed: DREADD occupancy and activation via converted clozapine. Science. 2017;357:503–507. doi: 10.1126/science.aan2475. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. González JC, Egea J, Del Carmen Godino M, Fernandez-Gomez FJ, Sánchez-Prieto J, Gandía L, García AG, Jordán J, Hernández-Guijo JM. Neuroprotectant minocycline depresses glutamatergic neurotransmission and Ca(2+) signalling in hippocampal neurons. Eur J Neurosci. 2007;26:2481–2495. doi: 10.1111/j.1460-9568.2007.05873.x. [DOI] [PubMed] [Google Scholar]
  13. Grace PM, Hutchinson MR, Manavis J, Somogyi AA, Rolan PE. A novel animal model of graded neuropathic pain: utility to investigate mechanisms of population heterogeneity. J Neurosci Methods. 2010;193:47–53. doi: 10.1016/j.jneumeth.2010.08.025. [DOI] [PubMed] [Google Scholar]
  14. Grace PM, Hutchinson MR, Maier SF, Watkins LR. Pathological pain and the neuroimmune interface. Nat Rev Immunol. 2014;14:217–231. doi: 10.1038/nri3621. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Grace PM, Strand KA, Galer EL, Urban DJ, Wang X, Baratta MV, Fabisiak TJ, Anderson ND, Cheng K, Greene LI, et al. Morphine paradoxically prolongs neuropathic pain in rats by amplifying spinal NLRP3 inflammasome activation. Proc Natl Acad Sci U S A. 2016a;113:E3441–3450. doi: 10.1073/pnas.1602070113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Grace PM, Gaudet AD, Staikopoulos V, Maier SF, Hutchinson MR, Salvemini D, Watkins LR. Nitroxidative Signaling Mechanisms in Pathological Pain. Trends Neurosci. 2016b;39:862–879. doi: 10.1016/j.tins.2016.10.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Gu N, Peng J, Murugan M, Wang X, Eyo UB, Sun D, Ren Y, DiCicco-Bloom E, Young W, Dong H, et al. Spinal Microgliosis Due to Resident Microglial Proliferation Is Required for Pain Hypersensitivity after Peripheral Nerve Injury. Cell Rep. 2016;16:605–614. doi: 10.1016/j.celrep.2016.06.018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Guan Z, Kuhn JA, Wang X, Colquitt B, Solorzano C, Vaman S, Guan AK, Evans-Reinsch Z, Braz J, Devor M, et al. Injured sensory neuron-derived CSF1 induces microglial proliferation and DAP12-dependent pain. Nat Neurosci. 2016;19:94–101. doi: 10.1038/nn.4189. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Harvey LO. Efficient estimation of sensory thresholds. Behav Res Methods Instrum Comput. 1986;18:623–632. [Google Scholar]
  20. Hayashi Y, Kawaji K, Sun L, Zhang X, Koyano K, Yokoyama T, Kohsaka S, Inoue K, Nakanishi H. Microglial Ca(2+)-activated K(+) channels are possible molecular targets for the analgesic effects of S-ketamine on neuropathic pain. J Neurosci Off J Soc Neurosci. 2011;31:17370–17382. doi: 10.1523/JNEUROSCI.4152-11.2011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Hoffmann A, Kann O, Ohlemeyer C, Hanisch UK, Kettenmann H. Elevation of basal intracellular calcium as a central element in the activation of brain macrophages (microglia): suppression of receptor-evoked calcium signaling and control of release function. J Neurosci Off J Soc Neurosci. 2003;23:4410–4419. doi: 10.1523/JNEUROSCI.23-11-04410.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Ji RR, Chamessian A, Zhang YQ. Pain regulation by non-neuronal cells and inflammation. Science. 2016;354:572–577. doi: 10.1126/science.aaf8924. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Lacagnina MJ, Watkins LR, Grace PM. Toll-like receptors and their role in persistent pain. Pharmacol Ther. 2017 doi: 10.1016/j.pharmthera.2017.10.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(−Delta Delta C(T)) Method. Methods San Diego Calif. 2001;25:402–408. doi: 10.1006/meth.2001.1262. [DOI] [PubMed] [Google Scholar]
  25. Milligan ED, Hinde JL, Mehmert KK, Maier SF, Watkins LR. A method for increasing the viability of the external portion of lumbar catheters placed in the spinal subarachnoid space of rats. J Neurosci Methods. 1999;90:81–86. doi: 10.1016/s0165-0270(99)00075-8. [DOI] [PubMed] [Google Scholar]
  26. Misko TP, Schilling RJ, Salvemini D, Moore WM, Currie MG. A fluorometric assay for the measurement of nitrite in biological samples. Anal Biochem. 1993;214:11–16. doi: 10.1006/abio.1993.1449. [DOI] [PubMed] [Google Scholar]
  27. Old EA, Clark AK, Malcangio M. The role of glia in the spinal cord in neuropathic and inflammatory pain. Handb Exp Pharmacol. 2015;227:145–170. doi: 10.1007/978-3-662-46450-2_8. [DOI] [PubMed] [Google Scholar]
  28. Parkhurst CN, Yang G, Ninan I, Savas JN, Yates JR, Lafaille JJ, Hempstead BL, Littman DR, Gan WB. Microglia promote learning-dependent synapse formation through brain-derived neurotrophic factor. Cell. 2013;155:1596–1609. doi: 10.1016/j.cell.2013.11.030. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Roth BL. DREADDs for Neuroscientists. Neuron. 2016;89:683–694. doi: 10.1016/j.neuron.2016.01.040. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Salter MW, Stevens B. Microglia emerge as central players in brain disease. Nat Med. 2017;23:1018–1027. doi: 10.1038/nm.4397. [DOI] [PubMed] [Google Scholar]
  31. Schreiber S, Getslev V, Backer MM, Weizman R, Pick CG. The atypical neuroleptics clozapine and olanzapine differ regarding their antinociceptive mechanisms and potency. Pharmacol Biochem Behav. 1999;64:75–80. doi: 10.1016/s0091-3057(99)00107-0. [DOI] [PubMed] [Google Scholar]
  32. Selfridge BR, Wang X, Zhang Y, Yin H, Grace PM, Watkins LR, Jacobson AE, Rice KC. Structure-Activity Relationships of (+)-Naltrexone-Inspired Toll-like Receptor 4 (TLR4) Antagonists. J Med Chem. 2015;58:5038–5052. doi: 10.1021/acs.jmedchem.5b00426. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Song ZP, Xiong BR, Guan XH, Cao F, Manyande A, Zhou YQ, Zheng H, Tian YK. Minocycline attenuates bone cancer pain in rats by inhibiting NF-κB in spinal astrocytes. Acta Pharmacol Sin. 2016;37:753–762. doi: 10.1038/aps.2016.1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Sorge RE, Mapplebeck JCS, Rosen S, Beggs S, Taves S, Alexander JK, Martin LJ, Austin JS, Sotocinal SG, Chen D, et al. Different immune cells mediate mechanical pain hypersensitivity in male and female mice. Nat Neurosci. 2015;18:1081–1083. doi: 10.1038/nn.4053. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Sweitzer SM, Hickey WF, Rutkowski MD, Pahl JL, DeLeo JA. Focal peripheral nerve injury induces leukocyte trafficking into the central nervous system: potential relationship to neuropathic pain. Pain. 2002;100:163–170. doi: 10.1016/s0304-3959(02)00257-9. [DOI] [PubMed] [Google Scholar]
  36. Szeto GL, Pomerantz JL, Graham DRM, Clements JE. Minocycline suppresses activation of nuclear factor of activated T cells 1 (NFAT1) in human CD4+ T cells. J Biol Chem. 2011;286:11275–11282. doi: 10.1074/jbc.M110.210518. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Tian DS, Peng J, Murugan M, Feng LJ, Liu JL, Eyo UB, Zhou LJ, Mogilevsky R, Wang W, Wu LJ. Chemokine CCL2-CCR2 Signaling Induces Neuronal Cell Death via STAT3 Activation and IL-1β Production after Status Epilepticus. J Neurosci Off J Soc Neurosci. 2017;37:7878–7892. doi: 10.1523/JNEUROSCI.0315-17.2017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Treutwein B, Strasburger H. Fitting the psychometric function. Percept Psychophys. 1999;61:87–106. doi: 10.3758/bf03211951. [DOI] [PubMed] [Google Scholar]
  39. Tsuda M, Shigemoto-Mogami Y, Koizumi S, Mizokoshi A, Kohsaka S, Salter MW, Inoue K. P2X4 receptors induced in spinal microglia gate tactile allodynia after nerve injury. Nature. 2003;424:778–783. doi: 10.1038/nature01786. [DOI] [PubMed] [Google Scholar]
  40. Wang X, Grace PM, Pham MN, Cheng K, Strand KA, Smith C, Li J, Watkins LR, Yin H. Rifampin inhibits Toll-like receptor 4 signaling by targeting myeloid differentiation protein 2 and attenuates neuropathic pain. FASEB J. 2013;27:2713–2722. doi: 10.1096/fj.12-222992. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Xu J, Dong H, Qian Q, Zhang X, Wang Y, Jin W, Qian Y. Astrocyte-derived CCL2 participates in surgery-induced cognitive dysfunction and neuroinflammation via evoking microglia activation. Behav Brain Res. 2017;332:145–153. doi: 10.1016/j.bbr.2017.05.066. [DOI] [PubMed] [Google Scholar]
  42. Yrjänheikki J, Keinänen R, Pellikka M, Hökfelt T, Koistinaho J. Tetracyclines inhibit microglial activation and are neuroprotective in global brain ischemia. Proc Natl Acad Sci U S A. 1998;95:15769–15774. doi: 10.1073/pnas.95.26.15769. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Zhang J, Shi XQ, Echeverry S, Mogil JS, De Koninck Y, Rivest S. Expression of CCR2 in both resident and bone marrow-derived microglia plays a critical role in neuropathic pain. J Neurosci Off J Soc Neurosci. 2007;27:12396–12406. doi: 10.1523/JNEUROSCI.3016-07.2007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Zhu H, Aryal DK, Olsen RHJ, Urban DJ, Swearingen A, Forbes S, Roth BL, Hochgeschwender U. Cre-dependent DREADD (Designer Receptors Exclusively Activated by Designer Drugs) mice. Genes N Y N 2000. 2016;54:439–446. doi: 10.1002/dvg.22949. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Zotova E, Bharambe V, Cheaveau M, Morgan W, Holmes C, Harris S, Neal JW, Love S, Nicoll JAR, Boche D. Inflammatory components in human Alzheimer’s disease and after active amyloid-β42 immunization. Brain. 2013;136:2677–2696. doi: 10.1093/brain/awt210. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

supplement. Figure S1. Effect of CNO on control BV-2 cells.

BV-2 cells were treated with a concentration range of CNO and a fixed concentration of LPS. CNO suppressed LPS-induced NO levels at higher concentrations, and did not affect cell viability.

NIHMS951471-supplement.pdf (27.9KB, pdf)

RESOURCES