Supplementary Information Development of small molecule inhibitors targeting adipose triglyceride lipase

Nicole Mayer 1#
Institute of Organic Chemistry
Graz University of Technology
8010GrazAustria

Martina Schweiger
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Matthias Romauch
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Gernot F Grabner
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Thomas O Eichmann
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Elisabeth Fuchs
Institute of Organic Chemistry
Graz University of Technology
8010GrazAustria

Jakov Ivkovic
Institute of Organic Chemistry
Graz University of Technology
8010GrazAustria

Christoph Heier
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Irina Mrak
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Achim Lass
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Gerald Höfler
Institute of Pathology
Medical University of Graz
8010GrazAustria

Christian Fledelius
Novo Nordisk A/S
Novo Nordisk ParkDK-2760MåløvDenmark

Rudolf Zechner
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Robert Zimmermann email@example.com
Institute of Molecular Biosciences
University of Graz
8010GrazAustria

Rolf Breinbauer firstname.lastname@example.com
Institute of Organic Chemistry
Graz University of Technology
8010GrazAustria

Supplementary Information Development of small molecule inhibitors targeting adipose triglyceride lipase
S-1 # These authors contributed equally to this work * Biological correspondence should be addressed to R. Zimmermann # Chemical correspondence should be addressed to R. Breinbauer Table of contents Supplementary Results Supplementary Note (Experimentals for Synthesis and Characterization of Compounds 1-5)
Supplementary Figures Supplementary Fig. 1. Dose-dependent inhibition of ATGLactivity in the presence of compound 3 and 4. The insert shows a Western blot confirming the expression of Strep-tagged proteins in E. coli at their correct molecular weight. The inhibitors were solubilized in DMSO and added at the indicated concentrations. DMSO alone was used as negative control. For determination of lipase activity, lysates were incubated with a substrate containing radiolabeled [9,10-3 H(N)]-triolein. Subsequently, FA were extracted and quantitated by liquid scintilation counting. Data are presented as mean ± S.D. of triplicate determinations and representative for at least three independent experiments.

S-2

Supplementary Results

Supplementary Table S1. Effect of Atglistatin on metabolic plasma parameters and tissue TG levels.

C57Bl6 mice were fasted over night and received Atglistatin by oral gavage or intraperitoneal injection (200 µmol/kg). After 8 h, blood was taken retro-orbitally and mice were sacrificed. Plasma TG, glucose, total cholesterol (TC), beta-hydroxybutyrate (beta-HB), and plasma insulin levels were measured using commercial assays. Data are presented as mean ± S.D. (n = 4-7; *, p < 0.05 **, p < 0.01).

Oral

Control + Atglistatin

Triglycerides (mmol/L) 0.58 ± 0.08 0.33 ± 0.14**   h after transfection, cells were harvested using a cell scraper and lysed in buffer C (50 mM Tris-Cl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1 % NP40) on ice. After centrifugation (1000 g, 4°C, 10 min), the supernatant (0.5 mg protein) was incubated with 10 µl Anti-Flag-Agarose over night at 4°C. After extensive washing in buffer C, the Flag-Agarose was heated to 99°C for 10 min in 2 x SDS sample buffer.

Thereafter, the input and the immunoprecipitated fraction were adjusted to SDS-PAGE. Flag-and Histagged proteins were detected using Flag-and His-antibodies.

S-14

Ethyl 1-(4-ethoxyphenyl)-1H-pyrazole-3-carboxylate (2)

Ethyl 1H-pyrazole-3-carboxylate (7)

A 100 mL Schlenk tube was dried under vacuum, filled with nitrogen and charged with 0.5 g

Ethyl 1-(4-ethoxyphenyl)-1H-pyrazole-3-carboxylate (2)

S-18

Ethyl 4`-ethoxybiphenyl-3-carboxylate (3)

S-19

A 20 mL Schlenk tube was dried under vacuum, filled with nitrogen and charged consecutively with 100.0 mg (70 µL, 0.437 mmol, 1.0 eq) ethyl-3-bromobenzoate, 72.5 mg (

3-(4'-(Dimethylamino)biphenyl-3-yl)-N,N-dimethylurea (4)

biphenyl)-4-amine (8)

A 100 mL Schlenk-flask, equipped with a magnetic stir bar and a stopper, was flame dried under vacuum, flushed with nitrogen and charged with 1.50 g (7.50 mmol, 1.0 eq) 4-bromo-N,Ndimethylaniline, 1.25 g (7.50 mmol, 1.0 eq) 3-nitrophenylboronic acid, 307.4 mg (0.38 mmol, S-20 0.05 eq) PdCl 2 (dppf)*DCM, 2.39 g (15.8 mmol, 2.1 eq) CsF and 60 mL anhydrous DME. The suspension was degassed by vacuum/N 2 cycles, heated to 80°C and stirred for 14 h. After full conversion according to TLC and GC-MS (filtration through a pipette filled with celite) the reaction mixture was cooled to room temperature and filtered through a pad of celite, which was rinsed with EtOAc. The solvent of the remaining filtrate was removed under reduced pressure.

The resulting dark brown solid was then purified via flash chromatography (CH/EtOAc = 19/1, size: 8.0 x 6.5 cm, 131 g silica gel) and dried under vacuum.

3-(4'-(Dimethylamino)biphenyl-3-yl)-N,N-dimethylurea (4)

A 100 mL Schlenk tube equipped with a magnetic stir bar and a stopper, was flame dried under vacuum, flushed with nitrogen and charged with 1.85 g (8.71 mmol, 1.0 eq) N 4' ,N 4' -dimethyl-

Fig. 2 .S- 5 SupplementaryFig. 3 .
253Toxicity test for Atglistatin in AML-12 mouse hepatocytes. For MTT-based in vitro viability assays, cells were seeded at an initial density of 1x10 4 cells per well in 96-well plates and cultured under standard conditions for twenty-four hours. The next day, cells were pretreated with different concentrations of Atglistatin dissolved in DMSO (A) or Cisplatin dissolved in dimethylformamide (DMF) (Sigma-Aldrich) as positive control for two hours (B). Medium was replaced by an identical fresh medium and incubated again for the indicated time points. Thereafter cells were incubated for 3 hours with 100 µl Thiazolyl Blue Tetrazolium Bromide (MTT). The resulting violet formazan crystals were dissolved by adding 100 µl of MTT Solubilization Solution (0.1% NP-40, 4 mM HCl and anhydrous isopropanol). After complete dissolution of the formazan product, absorbance was measured at 595 nm and 690 nm. The values obtained at 690 nm were subtracted from the 595 nm measurements. Measurements were performed in quadruplicates and are indicated as mean ± S.D. Data are representative for two independent experiments. *, p < 0.05 **, p < 0.01; ***, p < 0.001. Inhibition of ATGL in the absence (A) and in the presence of CGI-58 (B) by

Supplementary Fig. 8 .Fig. 9 .
89indicated concentrations. DMSO alone was used as negative control. (A) Effect of Atglistatin on murine MGL (purified from E. coli), and murine HSL activity (expressed in COS-7 cells). (B) Effect of Atglistatin on purified bovine LPL (Sigma Chemicals) and pancreatic lipase (from porcine pancreas, Sigma Chemicals). (C) Effect of Atglistatin on PNPLA6 and PNPLA7 activities (expressed in COS-7 cells). Data are presented as mean ± S.D. of triplicate determinations and are representative for at least two independent experiments (p < 0.05 **, p < 0.01; ***, p < 0.001). S-11 Supplementary Fig. 7. Inhibition of basal and stimulated lipolysis in 3T3-L1 adipocytes. Differentiated 3T3 cells were preincubated for 4 h with the indicated concentrations of Atglistatin. Thereafter, the medium was replaced by DMEM containing 2% BSA (FA free) with or without inhibitor. Medium was collected after incubation for another hour in the absence (A) and in the presence (B) of forskolin. Atglistatin was dissolved in DMSO and DMSO alone was used as negative control. Data are presented as mean ± S.D. (*, p < 0.05 **, p < 0.01; ***, p < 0.001). Effect of Atglistatin and the structural analogue 5 on ATGL activity and lipolysis. (A) Dose-dependent effect of Atglistatin and 5 on ATGL activity. Lysates from E. coli overexpressing ATGL and CGI-58 were incubated with a substrate containing radiolabeled [9,10-3 H(N)]triolein in the presence of the indicated concentrations of Atglistatin or 5 (B, C) Differentiated 3T3 cells were preincubated for 4 h in the presence of Atglistatin or 5. Thereafter, the medium was replaced by DMEM containing 2% BSA (FA free) and forskolin. Medium was collected after incubation for another hour in the presence or absence of Atglistatin and 5. Compounds were dissolved in DMSO and DMSO alone was used as negative control. Data are presented as mean ± S.D. (n=3; *, p < 0.05 **, p < 0.01; ***, p < 0.001). Time dependent (A) and dose-dependent (B) effect of an intraperitoneal injection of Atglistatin on plasma free fatty acids (FA) and glycerol levels of mice. C57Bl6 mice were fasted over night. For IP injection, the inhibitor was solubilized in PBS containing 0.25% Cremophor® EL by sonication. The time-dependent effect of Atglistatin injection was determined in the presence of 200µmol/kg Atglistatin. The dose-dependent effect was determined 8 h after inhibitor application. Blood was taken retroorbitally and plasma parameters were measured using commercial kits. Data are presented as mean ± S.D. (n = 5; * p < 0.05 **, p < 0.01; ***, p < 0.001).

1.1. 1 .. 2 .
12Ethyl 1-(4-ethoxyphenyl)-4-hydroxy-1H-pyrazole-3-carboxylate (1)S-22.214.171.124. Ethyl-4-chloro-2-((4-ethoxyphenyl)diazenyl)-3-oxobutanoate (6)A 10 mL one-neck round-bottom flask was charged with 250.0 mg (0.24 mL, 1.825 mmol, 1.04 eq) 4-ethoxyaniline which was dissolved in 3.1 mL acetic acid at 10 °C (ice bath). To this cooled solution 128.0 mg (1.861 mmol, 1.06 eq) NaNO 2 in 0.5 mL conc. H 2 SO 4 were added and the reaction mixture was stirred at 10 °C for 1 h. In a second 25 ml one-neck round-bottom flask 288.6 mg (0.24 mL, 1.755 mmol, 1.0 eq) ethyl-4-chloro-3-oxobutanoate were dissolved in a mixture of 1.3 mL acetic acid and 2.6 mL water and cooled to 0 °C (ice bath + NaCl). After one hour stirring at 10 °C the generated solution of the diazonium salt was given to the second solution at 0 °C and stirred for further 15 min at this temperature. An aqueous solution of 1.58 g (19.305 mmol, 11 eq) NaOAc in 3.0 mL water was added to the reaction mixture at 0 °C and the product precipitated. Stirring at rt over night, filtration, washing with a small amount of water and drying under high pressure yielded the crude product which was used in the next reaction step without further purification.yield: 453.7 mg (83 %); yellow-brown solid; M.p.: 106-110 °C; 1 H-NMR (300 MHz, DMSO-d 6 ): Isomer I (25 %): δ (ppm) = 14.39 (s, 1H, NH), 7.57-7.35 (m, 2H, Ar-H), 7.98-7.95 (m, 2H, Ar-H), 4.95-4.92 (m, 2H, Cl-CH 2 ), 4.31-4.27 (m, 2H, CH 2 ), 4.03-3.98 (m, 4H, 2 CH 2 ), 1.34-1.27 (m, 6H, 2 CH 3 ), Isomer II (75 %): δ (ppm) = 12.35 (s, 1H, NH), 7.57-7.35 (m, 2H, Ar-H), 6.98-6.95 (m, 2H, Ar-H), 4.95-4.92 (m, 2H, Cl-CH 2 ), 4.31-4.27 (m, 2H, CH 2 ), 4.03-3.98 (m, 4H, 2 CH 2 ), 1.34-1.27 (m, 6H, 2 CH 3 ); 13 C-NMR (75.5 MHz, DMSO-d 6 ): Isomer I: δ (ppm) = 187.6 (C=O), 164.0 (C=O), 156.9 (C q ), 135.2 (C=N), 125.1 (C q ), 118.1 (2 CH Ar ), 115.2 (2 CH Ar ), 63.2 (OCH 2 ), Ethyl 1-(4-ethoxyphenyl)-4-hydroxy-1H-pyrazole-3-carboxylate (1) A 25 mL Schlenk tube was charged with 270.6 mg (0.866 mmol, 1.0 eq) ethyl-4-chloro-2-((4ethoxyphenyl)diazenyl)-3-oxobutanoate (5) which was suspended in 4 mL EtOH under a gentle stream of nitrogen. After adding 102.0 mg (1.039 mmol, 1.2 eq) KOAc the suspension was stirred under reflux for 1.5 h, during which the suspension dissolved. TLC analysis (CH/EtOAc 3:1) indicated full conversion of the starting material. After cooling to rt the mixture was transferred to a flask to remove the EtOH with a rotary evaporator. The residue was dissolved in 20 mL EtOAc and washed with water (2 x 15 mL) and 15 mL brine, dried over MgSO 4 and concentrated at the rotary evaporator. Drying at high vacuum yielded the pure product without further purification. yield: 232.4 mg (97 %); dark red-brown solid; M.p.: 96-98 °C; R f (CH/ EtOAc 3:1): 0.35; 1 H-NMR (300 MHz, DMSO-d 6 ): δ (ppm) = 9.07 (s, 1H, OH), 7.96 (s, 1H, Ar-H), 7.70 (d, 3 J = 9.0Hz, 2H, Ar-H), 7.03 (d, 3 J = 9.0 Hz, 2H, Ar-H), 4.28 (q, 3 J = 6.9 Hz, 2H, CH 2 ), 4.06 (q, 3 J = 6.9Hz, 2H, CH 2 ), 1.36-1.27 (m, 6H, 2 CH 3 ); 13 C-NMR (75.5 MHz, DMSO-d 6 ): δ (ppm) = 161.6 (C=O), 157.3 (C q ), 145.0 (C q -OH), 132.8 (C q ), 131.1 (C q ), 120.0 (2 CH Ar ), 114.9 (2 CH Ar ), 114.8 (CH Ar ), 63.3 (CH 2 ), 59.7 (CH 2 ), 14.5 (CH 3 ), 14.2 (CH 3 ); GC-MS (NM_50_S2): t R = 8.071 min (m/z = 276.1, 99.0 % M + , BP).

( 4 . 1 (
41461 mmol, 1.0 eq) 1H-pyrazol-3-carboxylic acid which was dissolved in 20 mL EtOH. After adding 1.31 g (0.72 mL, 13.382 mmol, 3.0 eq) conc. H 2 SO 4 the colorless reaction mixture was heated to reflux (100 °C) and stirred at this temperature for 4 h. TLC analysis (DCM/MeOH 95:5) indicated full conversion of the starting material. After cooling to rt the mixture was transferred to a flask to remove the solvent at a rotary evaporator. The colorless residue was diluted in 20 mL water and neutralized with 17 mL saturated aqueous NaHCO 3 solution. Thereby a white solid precipitated. The aqueous layer was extracted with EtOAc (4 x 50 mL), dried over MgSO 4 and the solvent was evaporated under reduced pressure to yield the pure title compound.yield: 582.5 mg (93 %); colorless solid; M.p.: 158-161 °C; R f (DCM/MeOH 95:5): 0.42; 1 H-NMR (300 MHz, CDCl 3 ): δ (ppm) = 10.69 (bs, 1H, NH), 7.84 (d, 4 J = 2.1 Hz, 1H, Ar-H), 6.86 (d, 4 J = 2.1 Hz, 1H, Ar-H), 4.43 (q, 3 J = 7.2 Hz, 2H, CH 2 ), 1.41 (t, 3 J = 7.2 Hz, 3H, CH 3 ); 13 C-NMR (75.5 MHz, CDCl 3 ):δ (ppm) = 161.8 (C=O), 141.6 (C q ), 132.3 (CH Ar ), 107.8 (CH Ar ), 61.CH 2 ), 14.3 (CH 3 ); GC-MS (NM_50_S2): t R = 4.655 min (m/z = 140.1, 98.0 % M + , BP: 95.0).

p-bromophenetol and 1 mL dry ACN. The light brown suspension was degassed by vaccum/N 2 cycles and stirred first at 82 °C for 7 h and than after adding 0.5 mL dry DMF (solubility issue) at 120 °C for further 65 h. The GC-MS analysis showed full conversion. ACN and DMF were removed under high pressure and the brown residue was suspended in 10 mL EtOAc. After filtration of the brown suspension through a pad of silica and flushing with 150 mL EtOAc the colorless filtrate was concentrated under reduced pressure leading to 52.1 mg (39 %) crude product as a green-brown oil. Final purification by column chromatography (CH/EtOAc 3:1, size: 15.5 x 2.0 cm, 20 g silica gel) yielded the pure title compound. yield: 10.2 mg (8 %); orange solid; M.p.: 88-90 °C; R f (CH/ EtOAc 3:1): 0.40; 1 H-NMR (300 MHz, DMSO-d 6 ): δ (ppm) = 8.51 (d, 4 J = 2.4 Hz, 1H, Ar-H), 7.78 (d, 3 J = 9.0 Hz, 2H, Ar-H), 7.07 (d, 3 J = 9.0 Hz, 2H, Ar-H), 6.97 (d, 4 J = 2.4 Hz, 1H, Ar-H), 4.31 (q, 3 J = 6.9 Hz, 2H, CH 2 ), 4.08 (q, 3 J = 7.2 Hz, 2H, CH 2 ), 1.37-1.29 (m, 6H, 2 CH 3 ); 13 C-NMR (75.5 MHz, DMSO-d 6 ): δ (ppm) = 161.4 (C=O), 157.6 (C q ), 143.6 (C q ), 132.5 (C q ), 129.5 (CH Ar ), 120.7 (2 CH Ar ), 115.0 (2 CH Ar ), 119.8 (CH Ar ), 63.4 (CH 2 ), 60.3 (CH 2 ), 14.5 (CH 3 ), 14.1 (CH 3 ); GC-MS (NM_50_S2): t R = 7.722 min (m/z = 260.1, 98.0 % M + , BP).

dppf)*DCM and 5.0 mL DME. The orange suspension was degassed by vacuum/N 2 cycles and stirred at 80 °C for 7 h. GC-MS analysis indicated full conversion (98 % product) of the starting material. The reaction mixture was filtered through a pad of celite which was rinsed with EtOAc. The solvent from the filtrate was removed under reduced pressure and final purification by column chromatography (CH/EtOAc 15:1, size: 12.5 x 2.0 cm, 15 g silica gel) yielded the pure product. yield: 109.5 mg (93 %); colorless solid; M.p.: 46-47 °C; R f (CH/ EtOAc 15:1): 0.40; 1 H-NMR (300 MHz, CDCl 3 ): δ (ppm) = δ (ppm) = 8.25 (t, 4 J = 1.5 Hz, 1H, Ar-H), 7.98 (d, 3 J = 7.8 Hz, 1H, Ar-H), 7.74 (d, 3 J = 7.8 Hz, 1H, Ar-H), 7.56 (d, 3 J = 9.0 Hz, 2H, Ar-H), 7.48 (t, 3 J = 7.8 Hz, 1H, Ar-H), 6.98 (d, 3 J = 9.0 Hz, 2H, Ar-H), 4.41 (q, 3 J = 7.2 Hz, 2H, CH 2 ), 4.09 (q, 3 J = 6.9 Hz, 2H, CH 2 ), 1.47-1.39 (m, 6H, 2 CH 3 ); 13 C-NMR (75.5 MHz, CDCl 3 ): δ (ppm) = 166.6 (C=O), 158.8 (C q ), 141.0 (C q ), 132.5 (C q ), 131.0 (C q ), 130.9 (CH Ar ), 128.7 (CH Ar ), 128.2 (2 CH Ar ), 127.7 (CH Ar ), 127.6 (CH Ar ), 114.8 (2 CH Ar ), 63.8 (CH 2 ), 61.0 (CH 2 ), 14.8 (CH 3 ), 14.3 (CH 3 ); GC-MS (NM_50_S2): t R = 7.796 min (m/z = 270.1, 99 % M + , BP); HRMS (EI + ): m/z: calcd for C 17 H 18 O 3 [M] + : 270.1256; found 270.1260.

3 ); GC-MS (NM_50_S): t R = 8.171 min (m/z) = 242.0 (100 %, BP) [M + ], 226.0 (0.5 %) [M + -CH 3 ], 196.0 (23.9 %) [M + -NO 2 ], 180.0 (10.4 %) [M + -NO 2 , -CH 3 ], 152.0 (50.8 %) [M + -N(CH 3 ) 2 , -NO 2 ]; HRMS (M+H + ): m/z: calcd for C 14 H 14 O 2 N 2 [M+H] + : 243.281; found 243.2188.8.131.52.1 N 4' ,N 4' -Dimethyl-[1,1'-biphenyl]-3,4'-diamine (9)A solution of 2.21 g (9.13 mmol, 1.0 eq) N,N-dimethyl-3'-nitro-(1,1'-biphenyl)-4-amine in 92 mL THF was prepared to gain a deep orange 0.10 M solution of the substrate, which was then transferred into a 100 mL pear-shaped flask.For the reduction itself a continuous-flow hydrogenation reactor (Thales NANO H-Cube) with a 10 % Pd/C catalyst cartridge was used with the following conditions: 0.5 mL/min, 80°C, Full H 2 The product solution was collected in a 250 mL round-bottom flask. The solvent was removed under reduced pressure. The crude product was dried under vacuum and used for the next experiment without any further purification. yield: 2.054 g (quantitative yield); bright yellow solid; M.p.: 66-69 °C; R f (CH/ EtOAc 5:1): 0.10; 1 H-NMR (300 MHz, CDCl 3 ): δ (ppm) = 7.50 -7.47 (d, 2H, 3 J HH = 8.8 Hz, H-2/6), 7.22 -M+H + ): m/z: calcd for C 14 H 16 N 2 [M+H] + : 213.289; found 213.460.

: 2
21''/2''/4''/5''); GC-MS (NM_50_S): t R = 238.0 (100 %, BP) [M + -N(CH 3 ) 2 ], 195.0 (5.5 %) [M + -HN(CO)N(CH 3 ) 2 ], 180.0 (1.1 %) [M + -HN(CO)N(CH 3 ) 2 , -CH 3 ], 167.0 (4.1 %) [M + -1.5 N-(4'-(Dimethylamino)-[1,1'-biphenyl]-3-yl)-morpholine-4-carboxamide (5) Schlenk tube equipped with a magnetic stirring bar and a stopper, was flame dried under vacuum, flushed with nitrogen and charged with 206 mg (0.97 mmol, 1.0 eq) N 4' ,N 4' -dimethyl-[1,1'-biphenyl]-3,4'-diamine (9) and 2 mL anhydrous DCM. First 2.9 mg (23.6 µmol, 0.025 eq) DMAP, then 80 µL (79 mg, 0.99 mmol, 1.0 eq) anhydrous pyridine and 143 µL (183 mg, 1.22 mmol, 1.3 eq) 4-morpholinecarbonylchloride were added. When the yellow solution was heated up to 50°C a colour change to deep red was noticed. The reaction mixture was stirred for 2 d at 50°C and monitored using TLC and GC-MS (mini workup: dest. H 2 O, DCM, MgSO 4 ). After 2 d the GC-MS showed full conversion and the reaction was hydrolysed with 2 mL dest. H 2 O. The reaction mixture was transferred into a 100 mL roundbottom flask and the solvent was removed under reduced pressure. The red residue was dissolved in 20 mL DCM, dried over Na 2 SO 4 and the solvent was again removed under reduced pressure. The red crude product was then purified via flash chromatography (CH/EtOAc = 1/1, size: 14.8 x 3.0 cm, 40 g silica gel) and dried under vacuum. yield: 52.0 mg (0.17 mmol, 29 %), yellow solid. M.p.: 230-233 °C; R f (CH/EtOAc 1

Atglistatin. For determination of lipase activity, lysates of E. coli containing strep-tagged ATGL or ATGL and CGI-58 were incubated with the radiolabeled [9,10-3 H(N)]-triolein substrate in the presence of the indicated concentrations of Atglistatin. Subsequently, FA were extracted and quantitated by liquid scintillation counting. Lysates of cells expressing empty vector were set as blank. Measurements were performed in triplicates and are indicated as mean ± S.D. Data are representative for two independent Supplementary Fig. 4. LD localization of ATGL and the interaction of ATGL with CGI-58 is not affected by Atglistatin. (A) To determine whether Atglistatin displaces ATGL from lipid droplets, WAT lysates of wild-type mice were incubated in the presence or absence of 40 µM Atglistatin for 15 min at room temperature under constant shaking. Subsequently, lysates were fractionated in a sucrose gradient by ultracentrifugation for 1 h at 100,000 g. LDs were collected as a white band from the top of the tube and L1 cells and WAT lysates were subjected to Western blotting analysis. (C) To test whether Atglistatin disrupts the protein interaction of ATGL and CGI-58, Flag-tagged ATGL or empty Flag vector were cotransfected into HEK293T cells with either His-tagged LacZ or His-tagged CGI-58. 24experiments. *, p < 0.05 **, p < 0.01; ***, p < 0.001.

S-6

S-7

washed in buffer A by another centrifugation step for 1 h at 100,000 g. Total lysates, cytosolic fractions

(100.000 g supernatant), and lipid droplets were subjected to Western blotting analysis using antibodies

specific for murine ATGL (mATGL), perilipin, and GAPDH (Glycerinaldehyde 3-phosphate

dehydrogenase) and the respective secondary antibodies. WAT from ATGL-ko mice was used as negative

control. (B) To determine whether Atglistatin displaces ATGL from lipid droplets in living cells, 3T3-L1

adipocytes were incubated for 3 h in the presence or in the absence of 40 µM Atglistatin. Thereafter, cells

were disrupted and fractionated by differential centrifugation. Total fractions (TF), lipid droplets (LD) and

cytosol (Cyt) of 3T3-

Unless otherwise stated, all experiments were carried out under inert atmosphere by using standard Schlenk-techniques. Acetonitrile, DMF, Ethanol and Dimethoxyethane were purchased as absolute solvents from Acros Organics, Fisher Scientific and Sigma Aldrich. All applied starting materials were commercially available and were used as received: acetic acid (Acros, High pressure hydrogenation experiments were performed using the H-CubeTM continuous hydrogenation unit (HC-2.SS) from Thales Nanotechnology Inc. running with a Knauer Smartline pump 100 and equipped with a 10 mL ceramic pump head. As hydrogenation catalyst 10 % Pd/C catalyst cartridges were used (Thales Nanotechnology Inc., THS01111, 10% Pd/C CatCartTM). Silica gel chromatography was performed with Acros Organics silica gel 60 (35-70 µM). 1 H and 13 C NMR spectra were recorded on Bruker AVANCE III 300 spectrometer ( 1 H: 300.36 MHz; 13 C: 75.53 MHz) and chemical shifts are referenced to residual protonated solvent signals as internal standard. Electron impact (EI, 70eV) HRMS spectra were recorded on Waters GCT Premier equipped with direct insertion (DI) and GC (HP GC7890A). GC/MS analyses were made on an Agilent Technologies 5975 C inert MSD with Triple Axis Detector GC system with a mass sensitive detector.(Experimentals for Synthesis and Characterization of Compounds 1-5)

1. Experimental

1.1. General

96%), ethyl-4-chloro-3-oxobutanoate (Alfa Aesar, 97%), conc. sulphuric acid (Sigma Aldrich,

95-97%), 4-ethoxyaniline (Alfa Aesar, 98%), potassium acetate (Acros, 99+%), 1H-pyrazole-3-

carboxylic acid, (Aldrich, 97%), 3-nitrophenylboronic acid (Aldrich, 97%), ethyl-3-

bromobenzoate (Aldrich, 98%), 4-ethoxyphenylboronic acid (Aldrich), 4-bromo-N,N-

dimethylaniline (Aldrich, 97%), [1,1′-bis(diphenylphosphino)ferrocene]dichloropalladium(II)

complex
with
dichloromethane
(Aldrich),
cesium
fluoride
(Aldrich,
99%)

dimethylcarbamoylchloride (Aldrich, 98%), 4-morpholinecarbonyl chloride (Aldrich, 98 %).

[1,1'-biphenyl]-3,4'-diamine (2) and 25 mL anhydrous DCM. The resulting yellow solution was cooled to 0°C and first 1.21 mL (882 mg, 8.71 mmol, 1.0 eq) triethylamine and then 1.21 mL (1.41 g, 13.1 mmol, 1.5 eq) dimethylcarbamoyl chloride were added dropwise. The reaction wasmonitored using TLC and GC-MS (mini workup: dest. H 2 O, DCM, MgSO 4 ). After 5 d the GC-MS showed full conversion and the reaction was hydrolysed with 20 mL dest. H 2 O. The reaction mixture was transferred into a 250 mL round-bottom flask and the solvent was removed under reduced pressure. The brown residue was dissolved in 50 mL DCM, dried over Na 2 SO 4 and the solvent was removed under reduced pressure. The beige crude product was then purified via flash chromatography (CH/EtOAc = 1/1, size: 15 x 5.0 cm, 122 g silica gel) and dried undervacuum.

yield: 746 mg (30 %); beige solid; M.p.: 165-167 °C; R f (CH/ EtOAc 1:1): 0.29; 1 H-NMR

(300 MHz, CDCl 3 ): δ (ppm) = 7.59 (s, 1H, H-2'), 7.54 -7.51 (d, 2H, 3 J HH = 8.8 Hz,

H-4'/5'), 7.31 -7.30 (d, 2H, 3 J HH = 5.2 Hz, H-2/6), 7.23 -7.22 (d, 1H, 3 J HH = 5.8 Hz, H-6'), 6.89

-6.88 (d, 2H, 3 J HH = 6.0 Hz, H-3/5), 6.42 (s, 1H, NH), 3.04 (s, 6H, H-1''/2''), 3.0 (s, 6H, H-


Monosodium iodoacetate-induced joint pain is associated with increased phosphorylation of mitogen activated protein kinases in the rat spinal cord

Younglim Lee
Madhavi Pai
Jill-Desiree Brederson
Denise Wilcox
Gin Hsieh
Michael F Jarvis
Robert S Bitner
Monosodium iodoacetate-induced joint pain is associated with increased phosphorylation of mitogen activated protein kinases in the rat spinal cord
R E S E A R C H Open Access
Background: Intra-articular injection of monosodium iodoacetate (MIA) in the knee joint of rats disrupts chondrocyte metabolism resulting in cartilage degeneration and subsequent nociceptive behavior that has been described as a model of osteoarthritis (OA) pain. Central sensitization through activation of mitogen activated protein kinases (MAPKs) is recognized as a pathogenic mechanism in chronic pain. In the present studies, induction of central sensitization as indicated by spinal dorsal horn MAPK activation, specifically ERK and p38 phosphorylation, was assessed in the MIA-OA model. Results: Behaviorally, MIA-injected rats displayed reduced hind limb grip force 1, 2, and 3 weeks post-MIA treatment. In the same animals, activation of phospho ERK1/2 was gradually increased, reaching a significant level at post injection week 3. Conversely, phosphorylation of p38 MAPK was enhanced maximally at post injection week 1 and decreased, but remained elevated, thereafter. Double labeling from 3-wk MIA rats demonstrated spinal pERK1/2 expression in neurons, but not glia. In contrast, p-p38 was expressed by microglia and a subpopulation of neurons, but not astrocytes. Additionally, there was increased ipsilateral expression of microglia, but not astrocytes, in 3-wk MIA-OA rats. Consistent with increased MAPK immunoreactivity in the contralateral dorsal horn, mechanical allodynia to the contralateral hind-limb was observed 3-wk following MIA. Finally, intrathecal injection of the MEK1 inhibitor PD98059 blocked both reduced hind-limb grip force and pERK1/2 induction in MIA-OA rats. Conclusion: Results of these studies support the role of MAPK activation in the progression and maintenance of central sensitization in the MIA-OA experimental pain model.

Background

Osteoarthritis (OA), recognized as the most common form of degenerative arthritis, is caused by progressive disintegration of articular cartilage, bony overgrowth at the joint margins and synovial proliferation that can result in loss of joint function, disability and chronic pain [1][2][3]. The use of preclinical pain models to examine the pathogenic mechanisms responsible for OAinduced pain are being utilized for developing more effective therapeutic intervention [4,5]. A commonly used chemical model of OA pain involves intra-articular injection of the metabolic inhibitor monosodium iodoacetate (MIA) in the hind limb knee joint of rats, which disrupts chondrocyte glycolysis through inhibition of glyceraldehyde-3-phosphate dehydrogenase, leading to eventual cell death [6,7]. The progressive loss of chondrocytes following MIA results in histological and morphological changes of the articular cartilage similar to the pathology observed in OA patients [8][9][10]. In addition, focal bone damage observed with intra-articular MIA injection in rat has been reported to produce peripheral nerve injury as demonstrated by increased expression of the nerve injury marker ATF-3 (activating transcription factor-3) in L5 dorsal root ganglia, consistent with pathogenic changes associated with neuropathic pain [11].

However, analysis of pain behaviors such as weight bearing, tactile allodynia and mechanical hyperalgesia in the MIA-OA model have only recently been established, raising questions as to the appropriate behavioral endpoints for evaluating mechanisms and efficacy of novel analgesics for treating OA [4,7,12,13]. Determining biochemical signaling changes associated with nociceptive behaviors in MIA-injected animals may provide an alternative index of nociception, as well as improved understanding of cellular mechanisms involved in this model of OA pathology.

It has been demonstrated that during the first week following MIA injection, transient synovial inflammation may be the underlying cause of pain, whereas pain sensation in later stages may be due to biomechanical changes affecting the articular cartilage and subchondral bone [7]. Joint inflammation surrounding terminal endings of primary afferent neurons can be sensitized and activated by both normally innocuous and non-painful stimuli (peripheral sensitization) [14,15]. In turn, neurons in the spinal cord also become more responsive to innocuous and noxious stimuli onto the inflamed joint as well as adjacent non-inflamed normal tissue (central sensitization) [15,16]. Together, cellular sensitization in both peripheral and central sensory neurons is believed to be key in the initiation and maintenance of nociceptive transmission in chronic pain [17].

The causes leading to central sensitization of pain can be many-fold. It is known that primary afferent neurons release more transmitters upon stimulation following peripheral sensitization (presynaptic component), and neurons in the spinal cord are more excitable due to changes in receptor sensitivity (post synaptic component) [15]. One possible underling mechanisms for enhanced post synaptic sensitivity is up-regulation of second messenger system activation upon stimulation. Among various second messenger systems associated with pain responses, the family of mitogen-activated protein kinases (MAPKs) is likely candidates for development and maintenance of central pain sensitization. The MAPKs are serine/threonine protein kinases that include extracellular signal-regulated protein kinase (ERK) and p38 [18,19]. In the present study, we investigated the involvement of ERK1/2 and p38 phosphorylation-activation as an index of pain-induced central sensitization in the rat MIA model of osteoarthritis. Evaluating the temporal and activation profile of ERK1/2 and p38 may provide better understanding of disease progression in OA and the role of the MAPKs in development and maintenance of pain-induced central sensitization.

Results

MIA-induced pain behavior

Movement-induced pain behavior was measured using hind limb compressive grip force evaluation in which rats exhibit pain behaviors epitomized by a long-lasting decrement in bilateral compressive hind limb grip force following MIA-induced unilateral knee injury, as previously described [13]. Hind limb grip force was significantly reduced 1, 2, and 3 weeks following MIA injection into the hind-limb knee joint (Figure 1). At each time point, a similar reduction (~55%) in grip force was observed in all three OA groups as compared to non-pain controls (ANOVA; f = 101.7, p < 0.0001).

MIA-induced pERK1/2 immunoreactivity

Spinal cords were harvested (immediately following grip force testing) and immunohistochemically evaluated for changes in MAPK phosphorylation-activation at 1, 2, and 3-wk following intra-articular MIA injection. A significant overall increase in spinal pERK1/2 expression was observed in MIA-OA rats (ANOVA: F = 2.965, p = 0.0134), illustrated in Figure 2. Specifically, increased phospho-ERK1/2 immunoreactivity (black stained cells) was primarily observed in the upper lamina of the ipsilateral dorsal horn ( Figure 2B) that reached maximal levels at 3-wks as compared to naive (non-pain) controls ( Figure 2A). Similarly, a time-dependent increase in pERK1/2 expression was observed in the contralateral dorsal horn reaching maximal levels in the 3-wk MIA-OA group, albeit to a lesser extent in comparison to the ipsilateral side.

MIA-induced changes in p38 MAPK immunoreactivity

MIA-treated rats also displayed a significant increase in p38 phosphorylation-activation in the ipsilateral spinal dorsal horn ( Figure 3). However, in contrast to pERK,  Hind limb grip force following MIA knee injection in rat. MIA injection significantly decreased grip force 1, 2, and 3 wk post MIA injection. One-way ANOVA showed a significant treatment effect (F = 101.7, p < 0.0001); post-hoc analysis (Fisher's PLSD) revealed a significant grip force deficit in the 1, 2, and 3 wk-post MIA groups. Grip force strength was expressed as CFmax/kg body weight and represented as mean values (± S.E.M); ** p < 0.01 vs. naïve controls; n = 6/group. p38 phosphorylation was maximal at post injection week 1 and reduced thereafter (2 and 3 wk), although p-p38 expression remained significantly elevated compared to naïve controls at each time point ( Figure 3A). Cellular phospho-p38 immunoreactivity (black stained cells) was observed throughout the dorsal horn lamina ( Figure 3B). Similar to ERK, increased p-38 phosphorylation was observed in the contralateral dorsal horn, but to a lesser magnitude in comparison to ipsilateral side that was maximal in the 1-wk group and subsequently declining at 2 and 3 wk following MIA.

Cellular phenotype of spinal pERK1/2 and p-p38 expressing cells in MIA-OA rats

To determine the cellular phenotype of pERK1/2 and p-p38 expressing cells in the dorsal horn spinal cord of MIA-OA rats, double labeling experiments were conducted with the neuronal, astroglia, and microglia antibodies anti-NeuN, anti-GFAP and anti-OX-42, respectively ( Figure 4). Three weeks following MIA injection, ERK1/ 2 phosphorylation was observed in dorsal horn neurons as evidenced by the colocalization of anti-pERK1/2 and anti-NeuN immunoreactivity ( Figure 4A). In contrast, pERK1/2 expression was not observed in either astrocytes ( Figure 4B) or microglia ( Figure 4C). Conversely, p38 phosphorylation in the spinal dorsal horn was observed in microglia ( Figure 4F), but not astrocytes ( Figure 4E). In addition, spinal p-p38 expression was also observed in a subpopulation of small-diameter neurons ( Figure 4D

MIA-induced changes in OX-42 microglia and GFAP astroglia immunoreactivity

In addition to MAPK expression, spinal microglia activation was examined in OA rats 3-wk following MIA injection ( Figure 5). Increased expression of the microglia cell surface marker CD11b was observed in the ipsilateral, but not contralateral, dorsal horn 3-wk following MIA knee injection as compared to naïve (non pain) controls, as measured by OX-42 immunoreactivity ( Figure 5A). In contrast, there was no change in ipsilateral (or contralateral) astroglia expression in the dorsal horn 3-wk following MIA injection as compared to controls, as measured by GFAP immunoreactivity ( Figure 5B).

MIA-induced changes in pERK1/2 and mechanical allodynia nociceptive testing

To test whether increased MAPK activation observed in the contralateral (to MIA injection) dorsal horn translated to a nociceptive phenotype, mechanical allodynia was assessed on the contralateral paw 3-wk following MIA injection, as measuring grip force does not allow behavioral differentiation between the contra and ipsilateral paw ( Figure 6). Withdrawal threshold was significantly reduced (mechanical allodynia) in the contralateral paw 3-wk following MIA as compared to naïve control rats ( Figure 6A), a time when reduced grip strength is consistently observed [13]. In the same animals, elevated ERK1/2 phosphorylation was evident in both the ipsilateral and contralateral dorsal horn ( Figure 6B). The elevated pERK and mechanical allodynia observed in the contralateral spinal dorsal horn and paw, respectively, of MIA-OA rats supports biochemical translation to a nociceptive phenotype. pERK1/2 when immunohistochemically processed immediately following grip force testing, whereas PD98059-treated MIA-OA rats did not exhibit the same significant increase ( Figure 7B). Together, these results suggest that MIA-induced nociceptive behavior, i.e. reduced grip strength is associated with spinal pERK1/2 phosphorylation-activation.

Discussion

The use of intra-articular MIA as an animal model of OA has been previously reported to display multiple components of disease progression and symptoms akin to human OA pathology [8][9][10]. expressed in neurons, but not glia, was gradually increased following MIA injection and reached a significant level at post injection week 2 and 3 compared to naïve control. In contrast, enhanced phosphorylation of p38 MAPK, expressed primarily in microglia, was greatest at post injection week 1 and steadily reduced toward baseline thereafter. In addition, elevated MAPK phosphorylation was observed in the dorsal horn contralateral to the MIA-injected paw, which was accompanied by mechanical allodynia in the contralateral paw of 3-wk MIA-treated rats. Finally, it was demonstrated that i.t. administration of an ERK1/2 inhibitor in 3-wk MIA rats produced antinociception (normalized grip force strength) that correlated with attenuation of spinal pERK1/2 expression. Interestingly, spinal activation of microglia, but not astroglia, was also observed in MIAtreated rats.

It has been suggested that the early, transient synovial inflammation observed in MIA-treated rats may be the predominant cause of initial pain in MIA-OA rats, whereas later pain may result from biomechanical forces affecting the articular cartilage and subchondral bone [7]. It is interesting to speculate that different MAPKs may be involved in stages of OA disease progression, consistent with the temporal-dependent and differential profile of spinal ERK1/2 and p38 phosphorylation in MIA-OA rats observed in the present studies. Although the cellular mechanisms underlying chronic pain syndromes are not well understood, there is accumulating evidence supporting the role of plastic changes involving expression and function of ion channels, receptors and neurotransmitter-peptides in sensory systems responsible for pain transmission (e.g. spinal cord). Among the list of signaling molecules that may regulate the plasticity associated with chronic pain, MAPKs that include ERK and p38 have recently generated much interest [17,20]. As described here, the changes in MAPK phosphorylation-activation observed in MIA-injected rats, a novel finding, support a role of ERK1/2 and p38 in the development and maintenance of pain associated with OA pathology. Although we are not aware of prior reports examining MAPK expression in MIA-treated rats, or other experimental models of OA, neuropathic pain models involving spinal nerve injury have been well characterized for changes in MAPK phosphorylation involving central and peripheral sensitization. Specifically, activation of spinal ERK1/2 and p38 is induced following peripheral nerve injury in experimental models that includes: L5 spinal nerve ligation (SNL) and chronic constriction injury (CCI) of the sciatic nerve [21][22][23][24][25]. In general, studies conducted in nerve injury models have demonstrated that pERK1/2 activation occurs rapidly and transiently (10 min to 6 hr following nerve injury) in spinal dorsal horn neurons, with subsequent activation in glia cells, both microglia and astrocytes, two to 28 days later. In contrast, p38 induction appears to only occur in microglia, observed one to 14 days following nerve injury.

In the present studies, the diverse temporal profiles of spinal ERK1/2 and p38activation observed in MIA rats may reflect differential MAPK expression by distinct cell types, i.e. neurons and glia, as seen in nerve injury models. Specifically, expression of pERK1/2 was only observed in dorsal horn neurons at 3-wk following MIA, a time point where nociceptive behavior is well established and used in pharmacological antinociceptive testing [13]. In contrast, expression of p-p38 was primarily observed in microglia, but not astrocytes. However, a subpopulation of small diameter neurons in the superficial lamina, also expressed p-p38. It is interesting to note that microglia, but not astroglial, activation was observed in 3-wk MIA-treated rats, consistent with glial p-p38 expression restricted to microglia. Taken together, while there appears to be distinct temporal and biochemical discrepancies between our findings and pain models involving peripheral nerve injury, the pathogenic differences that likely exist between joint and nerve injury may explain these variations.

It is noteworthy that the contralateral spinal dorsal horn also showed a substantial increase in MAPK phosphorylation-activation following MIA injection. Moreover, mechanical allodynia was observed in the contralateral limb, demonstrating the parallel increase in MAPK activity has functional, i.e. pronociceptive consequences. In contrast, spinal MAPK activation reported in nerve injury models is primarily seen in the dorsal horn ipsilateral, but not contralateral, to injury [17]. Interestingly, there have been demonstrations of peripheral-nerve lesions that affect contralateral nonlesioned structures involving signaling via the system of commissural interneurons present in spinal cord and brainstem [26]. The excitatory communications between both sides of the spinal cord have been also demonstrated using electrophysiological techniques; Fitzgrald reported approximately 20% of cells in the substantia gelatinosa of the lumbar spinal cords showed a strong excitatory activation upon tetanic stimulation of the contra lateral sciatic nerve [27]. To our knowledge, the present data is the first demonstration of nociceptive-induced cellular signaling from ipsi to contralateral spinal dorsal horns following MIA injection, a finding that has not been observed in neuropathic peripheral nerve injury models such as SNL and CCI. However, Gao and colleagues recently demonstrated increased bilateral spinal cord expression of the MAPK JNK in the complete Freund's adjuvant model of persistent inflammatory pain [28]. Taken together, results of these studies and those presented here may suggest that the MIA-OA model share biochemical signaling properties of both neuropathic and inflammatory pain states.

It was observed that increased spinal ERK1/2 phosphorylation in 3-wk MIA-OA rats was blocked by the MEK inhibitor PD98059 when examined 30 min following acute intrathecal administration, as would be expected. Moreover, PD98059 treatment partially blocked the pain behavior, reduced grip force strength, observed in MIA-OA rats, supporting the potential involvement in part of ERK1/2 phosphorylation in the dorsal horn spinal cord in mediating nociceptiveinduced central sensitization associated with this model of OA. Consistent with our findings, the ability of PD98059 to produce antinociception has been previously demonstrated in various pain models including formalin [29], complete Freund's adjuvant-induced inflammation [30] and CCI [31].

Conclusions

Evaluation of biochemical pathways associated with central sensitization in animal models of OA has been lacking compared to neuropathic pain models involving peripheral nerve injury. Our current study demonstrates that activation of ERK1/2 and p38 MAPKs in the dorsal horn spinal cord is involved in nociceptive behaviors observed in the MIA-OA model. Furthermore, the ERK and p38 MAPK activation observed, which occurs primarily in neurons and microglia, respectively, displayed different temporal characteristics following MIA injection, suggesting possibly different roles of these MAPKs in development and maintenance of central pain sensitization. Taken together, these findings provide improved understanding of the biochemical relationship of MAPK activation and pain-induced central sensitization in the rat MIA-OA model, and may serve as a mechanistic tool for evaluating novel analgesic agents for the treatment of chronic pain associated with OA.

Methods

Animals

Adult male Sprague-Dawley rats (250-300 g, n = 6/ group) were used in experiments according to the internal Institutional Animal Care and Use Committee guidelines (Charles River Laboratories, Wilmington, MA). The animals were housed in Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC) approved facilities at Abbott Laboratories in a temperature-regulated environment under a controlled 12-h light-dark cycle, with lights on at 6:00 a. m. Food and water were available ad libitum at all times except during testing.

MIA injection: Osteoarthritic model of pain

Unilateral knee joint osteoarthritis was induced by a single intra articular (i.a.) injection of sodium monoiodoacetate (Sigma, St. Louis, MO) (MIA, 3 mg in 0.05 ml sterile isotonic saline) into the right knee joint cavity under light halothane (Halocarbon Laboratories, River Edge, NJ). Following injection, the animals were allowed to recover from the effects of anesthesia (usually 5-10 min) before returning them to their home cages.

To evaluate antinociceptive behavior and MAPK activation following MIA injection, separate groups of body weight matched Sprague-Dawley rats were injected with MIA on Day 0 (post MIA-injection week 3 group), Day 7 (post MIA-injection week 2 group), or Day 14 (post MIA-injection week 1 group). On Day 21, all MIAinjected animals as well as one group of naïve control animals were subjected to a grip force test. Twenty four hours later, all animals were perfused as described below.

MIA-induced nociceptive behavior at the contra lateral side was examined in a separate experiment. One group of animals was injected with MIA on Day 0 and allowed to recover for 21 days. On Day 21, the MIAinjected animals and one group of naïve control animals received grip force tests. On Day 22, a von Frey test was given to access contra lateral hind paw responses of all animals. Following 24 hour recovery, all animals were perfused, and spinal cords were harvested.

Intrathecal catheterization and injection of PD98059

The effects of a MEK inhibitor on MIA-induced pain behavior and pERK1/2 expression were evaluated. On the post MIA-injection Day 14, naive control animals, as well as MIA-injected animals received i.t. catheterization as previously described [32]. Briefly, rats were placed under isoflurane anesthesia and mounted onto a stereotaxic instrument using blunt ear bars, which held the animal's head firmly. An incision was made vertically from the dorsal surface of the occipital bone to the base of the skull (2 cm). Tissue was then displaced using a blunt probe so that the alanto-occipital membrane at the base of the skull was clearly seen. An intrathecal PE-10 catheter was inserted through the atlanto-occipital membrane via a small hole in the cisterna magnum. The catheter was then advanced 8.5 cm caudally such that the tip ended in the spinal subarachnoid space around the lumbar enlargement (L4-L6). The catheter was then secured to the musculature at the incision site. The incision was closed with surgical wound clips. The catheter was filled with sterile physiological saline and the ends of the catheter were heat-sealed. Animals with catheters were allowed 1 week of recovery from surgery before behavioral testing. The catheter was subsequently flushed with 10 μl of sterile water to maintain the patency.

On post MIA injection Day 21 (post IT catheterization Day 7), the animals were divided into four groups, one group of MIA-injected animals and one naïve control group animals were injected intrathecally with 10 μg (2 mg/10 ml/1 min/rat) of the MEK (ERK kinase) inhibitor PD98059 dissolved in a vehicle of 10% DMSO/ HBC (dimethyl sulfoxide/2-hydroxypropyl-β-cyclodextrin), while remaining groups were injected with the vehicle alone. Thirty minutes after i.t. injection, the animals were subjected to grip force test. Immediately after the behavioral test, the animals were perfused and their spinal cords were harvested.

Behavioral testing: Hind Limb Grip Force test

Measurements of peak hind limb grip force were conducted by recording the maximum compressive force exerted on the hind limb strain gauge setup, in a commercially available grip force measurement system (Columbus Instruments, Columbus, OH). During testing, each rat was gently restrained by grasping around its rib cage and then allowed to grasp the wire mesh frame (10-12 cm2) attached to the strain gauge. The experimenter then moved the animal in a rostral-tocaudal direction until the grip was broken. Each rat was sequentially tested twice at an approximately 2 min interval to obtain a raw mean grip force (CFmax). These raw mean grip force data were in turn converted to a maximum hind limb compressive force (CFmax) (gram force)/kg body weight for each animal. A group of agematched naïve animals was added and the data obtained from MIA-injected animals was compared to that of the naïve control group.

Behavioral testing: mechanical allodynia

Mechanical threshold was measured using calibrated von Frey monofilaments (Stoelting, Wood Dale, IL). Paw withdrawal threshold (PWT) was determined by increasing and decreasing stimulus intensity, and estimated using the Dixon's up-down method [33]. Rats were placed individually in inverted plastic containers (20 × 12.5 × 20 cm) on top of a suspended wire mesh with a 1 cm 2 grid to provide access to the ventral side of the hind paws. Rats were acclimated to the chambers for 20 min prior to testing. Monofilaments were presented perpendicularly to the plantar surface of the selected hind paw, and then held in this position for approximately 8 sec with enough force to cause a slight bend in the filament. Positive responses included an abrupt withdrawal of the hind paw from the stimulus, or flinching behavior immediately following removal of the stimulus. A 50% withdrawal threshold was determined using an up-down procedure [34].

Perfusion and tissue harvest

The animals were deeply anesthetized with CO 2 and perfused through the aorta with buffered saline (3 min, 25 ml/min) followed by 10% formalin (12 min, 30 ml/min). The spinal cords were extracted using hydraulic pressure and post-fixed in 10% formalin and stored in 20% sucrose PBS for overnight before sectioning.

Immunohistochemistry of free-floating spinal cord sections

After overnight incubation in 20% sucrose-PBS, the lumbar regions of spinal cords containing L3 to L5 were cut on a cryostat (40 μm coronal sections). Prior to sectioning, a knife cut was made through the ventral horn of spinal cords contra lateral to injection site for site specific evaluation of the data. The lumbar sections were immunostained in a free floating system for either anti-phospho p44/42 MAP Kinase (pERK 1/2) antibody (rabbit IgG, 1:700, Cell Signaling #9101), anti-phospho p38 MAP Kinase (Thr180/Tyr182) antibody (rabbit IgG, 1:1000, Cell Signaling # 9211L), anti-CD11b (OX-42) antibody (mouse IgG, 1:500, AbD Serotec), anti-GFAP antibody (mouse IgG, 1:1000, Millipore) using a threestep ABC-peroxidase technique beginning with a 30 min incubation with H 2 O 2 /PBS-Triton solution. Following PBS washes (3X); the sections were incubated with blocking serum (20 min) followed by the primary antibody for an overnight incubation. Twenty hrs later, the sections were washed with PBS (3x), and incubated for 1 hr with a secondary anti-rabbit IgG polyclonal antibody (goat IgG, 1:200; Vector Labs). Immunoreactivity was developed by an ABC (avidin-biotin)-peroxidase reaction using diaminobenzadine (DAB) as a chromogenic substrate. Sections were mounted, cover-slipped, and digitally photographed at 10× (Olympus BX-51 scope with F-View camera) with a shading correction to compensate for uneven illumination. Staining was quantified using an image analysis system (Olympus Micro-Suite Five) by measuring number of cells with homogeneous staining of a given antibody in laminae I-III of the spinal dorsal horns.

Double-labeling to determine the cellular phenotype of pERK1/2 and p-p38 expressing cells was carried out by antibody co-incubations of either anti-pERK1/2 or antip-p38 with the neuronal marker anti-NeuN, microglia marker anti-OX-42, or astroglia marker anti-GFAP. Specifically, 20 μm free floating lumbar spinal cord sections were blocked for 1-hr in 10% normal donkey serum, at room temperature. Sections were next coincubated with primary antiseras overnight at 4°C that consisted of anti-pERK (rabbit IgG, 1:500, Cell Signaling Technology) with either anti-NeuN (mouse IgG, 1:500, Millipore), anti-GFAP (mouse IgG, 1:1000, Millipore), or anti-OX-42 (mouse IgG, 1:500, AbD Serotec). Similar coincubations using the three cellular antibody markers were also made with anti-p-p38 (rabbit IgG, 1:500, Cell Signaling Technology). Following primary coincubations, sections were washed (3X, 2-hrs) in PBS and co-incubated with secondary fluorescence dye antibodies consisting of donkey-anti-rabbit Cy3 (1:500, Jackson Labs) and donkey anti-mouse Cy2 (1:500, Jackson Lab) for 1-hr at room temperature. Next, sections were washed (2X, 15 min) at room temperature, followed by an overnight wash in PBS at 4°C. Sections were then mounted on slides, dehydrated through series of ETOH (70, 95, 95, 100, 100%) for 2-3 minutes, then cleared in xylene and coverslipped with DPX. Cy2 (green,~492 nm excitation,~510 nm emission) and Cy3 (red,~550 nm excitation,~570 nm emission) fluorescence microscopy was conducted with an Olympus BX51 fluorescence microscope.

Statistics

Data were converted to percent change of ipsilateral naïve control and analyzed using one-way or two-way analysis of variance (ANOVA) followed by Fisher's protected least squares difference (PLSD) post-hoc analysis. A p-value < 0.05 was considered statistically significant.

Figure 1
1Figure 1 Hind limb grip force following MIA knee injection in rat. MIA injection significantly decreased grip force 1, 2, and 3 wk post MIA injection. One-way ANOVA showed a significant treatment effect (F = 101.7, p < 0.0001); post-hoc analysis (Fisher's PLSD) revealed a significant grip force deficit in the 1, 2, and 3 wk-post MIA groups. Grip force strength was expressed as CFmax/kg body weight and represented as mean values (± S.E.M); ** p < 0.01 vs. naïve controls; n = 6/group.

), in particular at the level of the superficial lamina(1)(2).

Figure 2
2ERK phosphorylation following MIA injection. ERK1/2 phosphorylation was increased in both ipsilateral and contralateral dorsal horn spinal cord at 2 and 3 wk following MIA compared to naïve control rats, (A). Phospho-ERK1/2 immunoreactivity (black stained cells) was observed in the upper lamina (hatched box) of the dorsal horn (B) that reached maximal levels at 3-wks as compared to naive (non-pain) controls. An overall ANOVA showed a significant main treatment effect (F = 2.965, p = 0.0134). All data are represented as mean values (± S.E.M) of the percent of ipsilateral naïve control rats; * p < 0.05, ** p < 0.01, vs. naïve controls (Fisher's PLSD post hoc analysis); n = 6/group.

Figure 3
3P38 phosphorylation following MIA injection. P38 phosphorylation was increased in both ipsilateral and contralateral dorsal horn spinal cord at 1, 2 and 3 wk following MIA as compared to naïve control rats (A). Phospho-p38 immunoreactivity (black stained cells) was observed throughout the upper lamina (hatched box) of the dorsal horn (B) that reached maximal levels at 1-wk and declined thereafter as compared to naive (non-pain) controls (ANOVA: F = 9.686, p < 0.0001). All data are represented as mean values (± S.E.M) of the percent of ipsilateral naïve control rats; * p < 0.05, ** p < 0.01 vs. naïve controls (Fisher's PLSD post hoc analysis); n = 5-7/group.

Figure 4
4MEK1 inhibitor, PD98059, on MIA-induced pain behavior and pERK1/2 expression To examine the functional role of spinal pERK in mediating nociceptive behavior, the MEK (ERK1/2 kinase) inhibitor PD98059 was tested in 3-wk MIA-OA rats. Intrathecal (i.t.) administration of PD98059 30-min before nociceptive behavior assessment significantly attenuated the MIA-induced reduction of grip force strength (Figure 7A). As expected, MIA-OA vehicle-i.t. controls rats displayed a significant increase in spinal Expression of spinal pERK1/2 and p-p38 in neuronal and glial cells of MIA-OA rats. Immunohistochemical double-labeling with the fluorescence dyes Cy2 (red; pERK1.2 or p-p38) and Cy3 (green; NeuN, GFAP, or OX-42) was conducted in lumbar spinal cord sections where orange-yellow fluorescence represents colocalization (white arrow). In the dorsal horn of OA rats 3-wk following MIA, double labeling with anti pERK1/2 (A-C) revealed colocalization with anti-NeuN/neuron (A), but not anti-GFAP/astrocyte (B) or anti-OX-42/microglia (C). Double labeling with anti-p-p38 (D-F) revealed colocalization with anti-NeuN (D), but limited to a subpopulation of small-diameter neurons. Colocalization of anti p-p38 in glial cells was not observed with anti-GFAP (E), but was observed with anti-OX-42 (F).

Figure 5
5However, demonstration of biochemical changes involving nociceptive signaling in this model are not as well established, in particular markers of central sensitization associated with chronic pain. The present study examined the development and maintenance MAPK phosphorylation-activation in the dorsal horn spinal cord as an index of central pain sensitization in the MIA-OA model. While MIA-injection into the hind limb joint reduced hind limb grip force asymptotically at all three time points tested (post injection week 1, 2, and 3), immunohistochemical evaluation of MAPK activation revealed differential temporal characteristics between pERK1/2 and phospho p38 MAPK. Specifically, pERK1/2 immunoreactivity in dorsal horn of spinal cord, Spinal glial activation in MIA-OA rats. Microglia, but not astrocyte, activation was observed in the ipsilateral dorsal horn 3-wk following MIA injection (Figure 5). In 3-wk MIA-treated rats, a significant increase in ipsilateral, but not contralateral, microglia expression was observed in the spinal dorsal horn, as determined by OX-42 (CD11b) immunoassaying, when compared to naive controls (A). In contrast, 3-wk MIA rats exhibited no differences in ipsilateral or contralateral astrocyte expression, as determined by GFAP immunoassaying, when compared to naive controls (B). All data are represented as mean values (± S.E.M) of the percent of ipsilateral naïve control rats; * p < 0.05 vs. naïve controls (Fisher's PLSD post hoc analysis); n = 6/group.

Figure 6 Figure 7
67Comparisons of MIA-induced contralateral paw allodynia to spinal ERK phosphorylation. Mechanical allodynia, measured as withdrawal threshold (***p < 0.001 vs. naïve control), was observed on the contralateral paw 3-wk following MIA (A). In the 3-wk MIA-treated rats, ERK phosphorylation was observed in both ipsilateral and contralateral dorsal horn spinal when immunohistochemically evaluated immediately following allodynia testing (B). ERK1/2 phosphorylation data are represented as mean values (± S.E.M) of the percent of ipsilateral naïve control rats; ** p < 0.01 vs. naïve controls (Fisher's PLSD post hoc analysis); Effects of MEK (ERK kinase) inhibitor PD98059 on pERK1/2 immunoreactivity in spinal dorsal horns. Intrathecal administration of PD98059 (10 μg) 30-min before nociceptive behavior assessment attenuated the MIA-induced reduction of grip force strength in 3-wk MIA rats (A). Grip force strength was expressed as CFmax/kg body weight; *p < 0.05, **p < 0.01 vs. naïve controls, # p < 0.05 vs. vehicle alone treated MIA rat; n = 6/group. MIA-OA vehicle-i.t. controls rats displayed a significant increase in spinal pERK1/2 when immunohistochemically processed immediately following grip force testing, whereas PD98059-treated MIA-OA rats did not exhibit the same significant increase (B). ERK phosphorylation data are represented as mean values (± S.E.M) of the percent of ipsilateral naïve control rats; ** p < 0.01 vs. naïve controls (Fisher's PLSD post hoc analysis); n = 6/group.

The strength of the maximum filament used for von Frey testing was 15 g. A percent maximal possible effect (% MPE) of testing compound was calculated according to the formula: ([compound -treated threshold] -[vehicle -treated threshold])/([maximum threshold] -[vehicle-treated threshold]) × 100%, where the maximum threshold was equal to 15 g.
Lee et al. Molecular Pain 2011, 7:39 http://www.molecularpain.com/content/7/1/39 Page 7 of 10
AcknowledgementsThe authors are grateful for the technical supports from many of our colleagues, particularly Art Nikkel.Authors' contributions YL conceived the study, participated in experimental design and interpretation, carried out immunohistochemical experiments and data analysis, and wrote the paper. MP carried out animal surgeries and behavioral pain testing, conducted data analysis for behavioral data, and wrote the paper. JDB carried out immunohistochemical experiments and data analysis. DW carried out immunohistochemical experiments and data analysis. GH participated in experimental design. MJ provided scientific advice for experimental design and manuscript preparation. RSB conceived the project, participated in experimental design and interpretation, and wrote the manuscript. All authors read and approved the text.Competing interestsThe authors declare that they have no competing interests.
Developments in the scientific understanding of osteoarthritis. S B Abramson, M Attur, Arthritis Res Ther. 11227Abramson SB, Attur M: Developments in the scientific understanding of osteoarthritis. Arthritis Res Ther 2009, 11:227.

Molecular mechanisms of cartilage destruction in osteoarthritis. R F Loeser, J Musculoskelet Neuronal Interact. 8Loeser RF: Molecular mechanisms of cartilage destruction in osteoarthritis. J Musculoskelet Neuronal Interact 2008, 8:303-6.

Osteoarthritic pain: a review of current, theoretical and emerging therapeutics. S J Read, A Dray, Expert Opin Investig Drugs. 17Read SJ, Dray A: Osteoarthritic pain: a review of current, theoretical and emerging therapeutics. Expert Opin Investig Drugs 2008, 17:619-40.

Pain related behaviour in two models of osteoarthritis in the rat knee. J Fernihough, C Gentry, M Malcangio, A Fox, J Rediske, T Pellas, B Kidd, S Bevan, J Winter, Pain. 112Fernihough J, Gentry C, Malcangio M, Fox A, Rediske J, Pellas T, Kidd B, Bevan S, Winter J: Pain related behaviour in two models of osteoarthritis in the rat knee. Pain 2004, 112:83-93.

Estimates of the prevalence of arthritis and selected musculoskeletal disorders in the United States. R C Lawrence, C G Helmick, F C Arnett, R A Deyo, D T Felson, E H Giannini, S P Heyse, R Hirsch, M C Hochberg, G G Hunder, M H Liang, S R Pillemer, V D Steen, F Wolfe, Arthritis Rheum. 41Lawrence RC, Helmick CG, Arnett FC, Deyo RA, Felson DT, Giannini EH, Heyse SP, Hirsch R, Hochberg MC, Hunder GG, Liang MH, Pillemer SR, Steen VD, Wolfe F: Estimates of the prevalence of arthritis and selected musculoskeletal disorders in the United States. Arthritis Rheum 1998, 41:778-99.

Monoiodoacetate-induced experimental osteoarthritis: a dose-response study of loss of mobility, morphology, and biochemistry. C Guingamp, P Gegout-Pottie, Philippe L Terlain, B Netter, P Gillet, P , Arthritis Rheum. 40Guingamp C, Gegout-Pottie P, Philippe L, Terlain B, Netter P, Gillet P: Mono- iodoacetate-induced experimental osteoarthritis: a dose-response study of loss of mobility, morphology, and biochemistry. Arthritis Rheum 1997, 40:1670-9.

Antinociceptive efficacy of lacosamide in the monosodium iodoacetate rat model for osteoarthritis pain. B Beyreuther, N Callizot, T Stohr, Arthritis Res Ther. 914Beyreuther B, Callizot N, Stohr T: Antinociceptive efficacy of lacosamide in the monosodium iodoacetate rat model for osteoarthritis pain. Arthritis Res Ther 2007, 9:R14.

Animal models of osteoarthritis. A M Bendele, J Musculoskelet Neuronal Interact. 1Bendele AM: Animal models of osteoarthritis. J Musculoskelet Neuronal Interact 2001, 1:363-76.

The monosodium iodoacetate model of osteoarthritis: a model of chronic nociceptive pain in rats?. R Combe, S Bramwell, M J Field, Neurosci Lett. 370Combe R, Bramwell S, Field MJ: The monosodium iodoacetate model of osteoarthritis: a model of chronic nociceptive pain in rats? Neurosci Lett 2004, 370:236-40.

Sodium iodoacetate-induced experimental osteoarthritis and associated pain model in rats. K Kobayashi, R Imaizumi, H Sumichika, H Tanaka, M Goda, A Fukunari, H Komatsu, J Vet Med Sci. 65Kobayashi K, Imaizumi R, Sumichika H, Tanaka H, Goda M, Fukunari A, Komatsu H: Sodium iodoacetate-induced experimental osteoarthritis and associated pain model in rats. J Vet Med Sci 2003, 65:1195-9.

Structural pathology in a rodent model of osteoarthritis is associated with neuropathic pain: increased expression of ATF-3 and pharmacological characterisation. S P Ivanavicius, A D Ball, C G Heapy, F R Westwood, F Murray, S J Read, Pain. 128Ivanavicius SP, Ball AD, Heapy CG, Westwood FR, Murray F, Read SJ: Structural pathology in a rodent model of osteoarthritis is associated with neuropathic pain: increased expression of ATF-3 and pharmacological characterisation. Pain 2007, 128:272-82.

Weight bearing as a measure of disease progression and efficacy of anti-inflammatory compounds in a model of monosodium iodoacetate-induced osteoarthritis. S E Bove, S L Calcaterra, R M Brooker, C M Huber, R E Guzman, P L Juneau, D J Schrier, K S Kilgore, Osteoarthritis Cartilage. 11Bove SE, Calcaterra SL, Brooker RM, Huber CM, Guzman RE, Juneau PL, Schrier DJ, Kilgore KS: Weight bearing as a measure of disease progression and efficacy of anti-inflammatory compounds in a model of monosodium iodoacetate-induced osteoarthritis. Osteoarthritis Cartilage 2003, 11:821-30.

Pharmacological modulation of movement-evoked pain in a rat model of osteoarthritis. P Chandran, M Pai, E A Blomme, G C Hsieh, M W Decker, P Honore, Eur J Pharmacol. 613Chandran P, Pai M, Blomme EA, Hsieh GC, Decker MW, Honore P: Pharmacological modulation of movement-evoked pain in a rat model of osteoarthritis. Eur J Pharmacol 2009, 613:39-45.

How do we manage chronic pain?. H G Schaible, H Vanegas, Baillieres Best Pract Res Clin Rheumatol. 14Schaible HG, Vanegas H: How do we manage chronic pain? Baillieres Best Pract Res Clin Rheumatol 2000, 14:797-811.

Von Banchet GS: Mechanisms of pain in arthritis. H G Schaible, A Ebersberger, Ann N Y Acad Sci. 966Schaible HG, Ebersberger A, Von Banchet GS: Mechanisms of pain in arthritis. Ann N Y Acad Sci 2002, 966:343-54.

Afferent and spinal mechanisms of joint pain. H G Schaible, B D Grubb, Pain. 55Schaible HG, Grubb BD: Afferent and spinal mechanisms of joint pain. Pain 1993, 55:5-54.

MAP kinase and pain. R R Ji, Gereau Rwt, M Malcangio, G R Strichartz, Brain Res Rev. 60Ji RR, Gereau RWt, Malcangio M, Strichartz GR: MAP kinase and pain. Brain Res Rev 2009, 60:135-48.

The MAPK signaling cascade. R Seger, E G Krebs, Faseb J. 9Seger R, Krebs EG: The MAPK signaling cascade. Faseb J 1995, 9:726-35.

Mitogen-activated protein kinase: conservation of a three-kinase module from yeast to human. C Widmann, S Gibson, M B Jarpe, G L Johnson, Physiol Rev. 79Widmann C, Gibson S, Jarpe MB, Johnson GL: Mitogen-activated protein kinase: conservation of a three-kinase module from yeast to human. Physiol Rev 1999, 79:143-80.

Intracellular signaling in primary sensory neurons and persistent pain. J K Cheng, R R Ji, Neurochem Res. 33Cheng JK, Ji RR: Intracellular signaling in primary sensory neurons and persistent pain. Neurochem Res 2008, 33:1970-8.

ERK is sequentially activated in neurons, microglia, and astrocytes by spinal nerve ligation and contributes to mechanical allodynia in this neuropathic pain model. Z Y Zhuang, P Gerner, C J Woolf, R R Ji, Pain. 114Zhuang ZY, Gerner P, Woolf CJ, Ji RR: ERK is sequentially activated in neurons, microglia, and astrocytes by spinal nerve ligation and contributes to mechanical allodynia in this neuropathic pain model. Pain 2005, 114:149-59.

Partial sciatic nerve ligation induces increase in the phosphorylation of extracellular signal-regulated kinase (ERK) and c-Jun N-terminal kinase (JNK) in astrocytes in the lumbar spinal dorsal horn and the gracile nucleus. W Ma, R Quirion, Pain. 99Ma W, Quirion R: Partial sciatic nerve ligation induces increase in the phosphorylation of extracellular signal-regulated kinase (ERK) and c-Jun N-terminal kinase (JNK) in astrocytes in the lumbar spinal dorsal horn and the gracile nucleus. Pain 2002, 99:175-84.

Identification of MEK1 as a novel target for the treatment of neuropathic pain. A Ciruela, A K Dixon, S Bramwell, M I Gonzalez, R D Pinnock, K Lee, Br J Pharmacol. 138Ciruela A, Dixon AK, Bramwell S, Gonzalez MI, Pinnock RD, Lee K: Identification of MEK1 as a novel target for the treatment of neuropathic pain. Br J Pharmacol 2003, 138:751-6.

p38 mitogen-activated protein kinase is activated after a spinal nerve ligation in spinal cord microglia and dorsal root ganglion neurons and contributes to the generation of neuropathic pain. Sx Jin, Z Y Zhuang, C J Woolf, Ji Rr, J Neurosci. 23SX Jin, Zhuang ZY, Woolf CJ, RR Ji: p38 mitogen-activated protein kinase is activated after a spinal nerve ligation in spinal cord microglia and dorsal root ganglion neurons and contributes to the generation of neuropathic pain. J Neurosci 2003, 23:4017-22.

Activation of p38 mitogen-activated protein kinase in spinal hyperactive microglia contributes to pain hypersensitivity following peripheral nerve injury. M Tsuda, A Mizokoshi, Y Shigemoto-Mogami, S Koizumi, K Inoue, Glia. 45Tsuda M, Mizokoshi A, Shigemoto-Mogami Y, Koizumi S, Inoue K: Activation of p38 mitogen-activated protein kinase in spinal hyperactive microglia contributes to pain hypersensitivity following peripheral nerve injury. Glia 2004, 45:89-95.

Does the right side know what the left is doing?. M Koltzenburg, P D Wall, S B Mcmahon, Trends Neurosci. 22Koltzenburg M, Wall PD, McMahon SB: Does the right side know what the left is doing? Trends Neurosci 1999, 22:122-7.

Influences of contralateral nerve and skin stimulation on neurones in the substantia gelatinosa of the rat spinal cord. M Fitzgerald, Neurosci Lett. 36Fitzgerald M: Influences of contralateral nerve and skin stimulation on neurones in the substantia gelatinosa of the rat spinal cord. Neurosci Lett 1983, 36:139-43.

The c-Jun N-terminal kinase 1 (JNK1) in spinal astrocytes is required for the maintenance of bilateral mechanical allodynia under a persistent inflammatory pain condition. Y J Gao, Z Z Xu, Y C Liu, Y R Wen, I Decosterd, R R Ji, Pain. 148Gao YJ, Xu ZZ, Liu YC, Wen YR, Decosterd I, Ji RR: The c-Jun N-terminal kinase 1 (JNK1) in spinal astrocytes is required for the maintenance of bilateral mechanical allodynia under a persistent inflammatory pain condition. Pain 2010, 148:309-19.

Involvement of phosphorylated Ca2+/calmodulin-dependent protein kinase II and phosphorylated extracellular signal-regulated protein in the mouse formalin pain model. S S Choi, Y J Seo, E J Shim, M S Kwon, J Y Lee, Y O Ham, H W Suh, Brain Res. 1108Choi SS, Seo YJ, Shim EJ, Kwon MS, Lee JY, Ham YO, Suh HW: Involvement of phosphorylated Ca2+/calmodulin-dependent protein kinase II and phosphorylated extracellular signal-regulated protein in the mouse formalin pain model. Brain Res 2006, 1108:28-38.

Inhibition of ERK phosphorylation decreases nociceptive behaviour in monoarthritic rats. C D Cruz, F L Neto, J Castro-Lopes, S B Mcmahon, F Cruz, Pain. 116Cruz CD, Neto FL, Castro-Lopes J, McMahon SB, Cruz F: Inhibition of ERK phosphorylation decreases nociceptive behaviour in monoarthritic rats. Pain 2005, 116:411-9.

Intracisternal administration of mitogen-activated protein kinase inhibitors reduced mechanical allodynia following chronic constriction injury of infraorbital nerve in rats. E J Lim, H J Jeon, G Y Yang, M K Lee, J S Ju, S R Han, D K Ahn, Prog Neuropsychopharmacol Biol Psychiatry. 31Lim EJ, Jeon HJ, Yang GY, Lee MK, Ju JS, Han SR, Ahn DK: Intracisternal administration of mitogen-activated protein kinase inhibitors reduced mechanical allodynia following chronic constriction injury of infraorbital nerve in rats. Prog Neuropsychopharmacol Biol Psychiatry 2007, 31:1322-9.

Chronic catheterization of the spinal subarachnoid space. T L Yaksh, T A Rudy, Physiol Behav. 17Yaksh TL, Rudy TA: Chronic catheterization of the spinal subarachnoid space. Physiol Behav 1976, 17:1031-6.

Efficient analysis of experimental observations. W J Dixon, Annu Rev Pharmacol Toxicol. 20Dixon WJ: Efficient analysis of experimental observations. Annu Rev Pharmacol Toxicol 1980, 20:441-62.

Quantitative assessment of tactile allodynia in the rat paw. S R Chaplan, F W Bach, J W Pogrel, J M Chung, T L Yaksh, 10.1186/1744-8069-7-39J Neurosci Methods. 53Chaplan 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:10.1186/1744-8069-7-39

Monosodium iodoacetate-induced joint pain is associated with increased phosphorylation of mitogen activated protein kinases in the rat spinal cord. Lee, Molecular Pain. 39Cite this article asCite this article as: Lee et al.: Monosodium iodoacetate-induced joint pain is associated with increased phosphorylation of mitogen activated protein kinases in the rat spinal cord. Molecular Pain 2011 7:39.

. Lee, Molecular Pain. 739Lee et al. Molecular Pain 2011, 7:39