Adult Sprague–Dawley rats (n = 135, 6–8 weeks, 180–220 g) were obtained from the Experimental Animal Center of Tianjin Medical University Cancer Institute and Hospital (Tianjin, China) and Beijing Vital River Laboratory Animal Technology Co., Ltd., and housed under controlled relative humidity (20–30%), temperature (23 ± 2°C), and 12/12 h light/dark cycle (light from 08:00 to 20:00) with access to food and water ad libitum. Before experimental procedures, the animals were allowed 7 days of acclimation, and efforts were made to limit any distress. The protocol was approved by the Animal Care and Use Committee of the Animal Ethical and Welfare Committee of Tianjin Medical University Cancer Institute and Hospital (approval #NSFC-AE-2020210). The study was conducted in accordance with the Declaration of the National Institutes of Health’s Guide for the Care and Use of Laboratory Animal (Publication No. 80-23, revised 1996) and the policies on the use of laboratory animals issued by the International Association for the Study of Pain (IASP).

The MTL enzyme-linked immunosorbent assay (ELISA) kit was purchased from Gene Biotech Company (Shanghai, China). The doses of each drug were determined based on preliminary experiments. The MTL (NBP2-93868; Novus Biologicals, Littleton, CO, USA), MTLR (DF4993; Affinity BIO, Scoresby, Australia), IgG-FITC (Santa Cruz Biotechnology, Santa Cruz, CA, USA), IgG-CY3 (Santa Cruz Biotechnology, Santa Cruz, CA, USA), IgG-CY5 (Santa Cruz Biotechnology, Santa Cruz, CA, USA), and HRP-conjugated (GB23303; Servicebio, Woburn, MA, USA) antibodies were purchased.

Seventy-eight rats were randomly divided into two groups: the control group (group C, n = 6) and the incisional pain group (group P, n = 72). Group P underwent incisional pain modeling. Six rats in the two groups were randomly selected. The mechanical withdrawal threshold (MWT) and thermal withdrawal latency (TWL) were measured 24 h before operation (T0), 2 h after operation (T1), 24 h after operation (T2), 48 h after operation (T3), and 72 h after operation (T4).

In order to demonstrate the effect of MTL on acute pain, we performed direct intrathecal injections of different doses of MTL in rats. Forty-five healthy male SD rats (6–8 weeks, 180–220 g) were randomized into five groups (n = 9/group): control group (group C), incisional pain group (group P), incisional pain + 1 μg MTL group (group 1MP), incisional pain + 5 μg MTL group (group 5MP), and incisional pain + 25 μg MTL group (group 25MP). The MWT and TWL were measured at T0, T1, T2, T3, and T4.

To measure locomotor function, we performed open field testing in which 12 SD rats (6–8 weeks, 180–220 g) were randomized into two groups: saline group (group Vehicle, n = 6) and intrathecal MTL group (group MTL, n = 6). 25 μl of normal saline was injected intrathecally in group Vehicle and 25 μg of MTL in 25 μl saline was injected in group MTL. Locomotor behavior was recorded 2 h after injection.

A lumbar intrathecal catheter was inserted using a modified version of a reported technique for subarachnoid drug administration (Storkson et al., 1996). Briefly, rats were anesthetized with 4% chloral hydrate (400 mg/kg, intraperitoneal injection). The lumbar skin was shaved, cleaned, and incised. The intervertebral space between L5 and L6 was punctured with a hypodermic needle, and a PE10 tubing (outer diameter of 0.55 mm, inner diameter of 0.30 mm; AniLab Software and Instruments Company, China) was inserted. The catheter localization was verified by injecting 10 μl of 2% lidocaine: only the rats with a brief bilateral hind limb paresis after injection (indicated catheterization success) were used for the incisional pain model and intrathecal MTL injection experiments (Ray et al., 2011; Hou et al., 2016). The experiments started 7 days after successful catheter placement. At 20 min before modeling, 25 μl of normal saline was injected intrathecally in groups C and P, while the 1MP, 5MP, and 25MP were injected with the proper dose of MTL in 25 μl of normal saline. After 20 min, the right plantaris muscle incision was made in the pain groups.

The rat hind paw plantar incisional pain model was established as previously described (Brennan et al., 1996). After the rats were anesthetized using sevoflurane, a 1 cm longitudinal incision was made through the skin and fascia of the plantar aspect of the right hind paw. The underlying plantar muscle was elevated and incised longitudinally, leaving the muscle origin and insertion intact. The skin was apposed with two mattress sutures using 5-0 nylon threads. After surgery, the animals were allowed to recover in their cages.

The MWT was determined as previously described (Hohmann et al., 2017). In order to determine the MWT of the rats, a dynamic plantar esthesiometer (Ugo Basile, Comerio, Italy) was used. The rats were put on an elevated grid in test cages at least 1 h before the measurement to allow accommodation. Then, baseline measurements were performed before incisional pain modeling. A steel filament was pushed against the plantar side of the hind paw with a linear ascending force (0–5 g over 10 s, in 0.5 g/s intervals) until a fast withdrawal of the paw was observed. The time between the first contact of the steel rod and the paw withdrawal was measured in seconds (paw withdrawal latency, PWL), and a cut-off time of 20 s was set.

The TWL was determined as previously described (Cao et al., 2015). TWL was evaluated using the Plantar Test apparatus (UgoBasile, Comerio, Italy). The Plantar Test apparatus was calibrated before TWL measurement. Each animal was allowed to habituate to the environment for 1 h before any measurements. A mobile heat source with an infrared intensity of 70 W was placed under the hind paw. TWL was defined as the duration from starting heat to inducing the paw to withdraw. Each foot was measured three times, and then its average was calculated. There was a 5-min rest period between measurements.

Motilin levels in the plasma, stomach, duodenum, and spinal cord were determined using rabbit-specific ELISA kits per the manufacturer’s instructions. Six rats in each group were sacrificed by spine dislocation before incision pain at T0–T4. The plasma, stomach, duodenum, and spinal cord samples were quickly prepared for MTL assessment.

The spinal cord samples from six rats at each time point were quickly extracted and stored in liquid nitrogen. Tissue samples were homogenized at pH 7.4 in lysis buffer containing Tris (20.0 mM), sucrose (250.0 mM), sodium orthovanadate (0.03 mM), magnesium chloride (2.0 mM), ethylenediaminetetraacetic acid (2.0 mM), ethylene glycol tetraacetic acid (2.0 mM), phenylmethylsulfonyl fluoride (2.0 mM), dithiothreitol (1.0 mM), and a protease inhibitor cocktail (0.02% v/v; G2006, Servicebio, Wuhan, China). The homogenates were centrifuged at 5,000 × g for 30 min at 4°C. The supernatants were stored at –80°C. Protein concentration was measured using the Bradford method. Proteins (30 μg) were separated by 10% SDS-PAGE and transferred onto a nitrocellulose membrane (G6014, Servicebio, Wuhan, China). The membranes were incubated overnight at 4°C with primary polyclonal rabbit anti-human MTLR IgG (1:1,000; DF4993; Affinity biosciences, USA) and Motilin Antibody (1:1,000; NBP2-93868; Novus Biologicals, Littleton, CO, USA). After washing with Tris-buffered saline Tween 20 (TBST), the membrane was incubated for 1 h with secondary antibody conjugated with alkaline phosphatase (1:500) at room temperature. The immune complexes were detected using a nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate assay kit (Sigma, St. Louis, MO, USA). Western blot densitometry analysis of signal intensity was performed using Adobe Photoshop software (Adobe, San Jose, CA, USA). The blot density from control groups was set at 100%.

At T0, T1, and T2, the rats were anesthetized with 10% chloral hydrate (300 mg/kg, i.p.) and perfused intracardially with 200 ml of saline followed by 500 ml of ice-cold 4% paraformaldehyde in 0.2 M phosphate buffer (pH 7.4). The lumbar spinal cord (at L4-6) was removed immediately, postfixed in the same fixative overnight, and subsequently transferred to 30% sucrose (w/v) in phosphate buffer and stored at 4°C for several days. The specimens were then embedded with OCT at –20°C, and 30-μm transverse series sections were cut on a cryostat (CM1900, Leica, Wetzlar, Germany) and collected in 0.01 M phosphate-buffered saline (PBS, pH 7.4).

The sections were immersed in antigen retrieval buffer, heated in a microwave oven for antigen retrieval at a subboiling temperature, cooled naturally, and washed with PBS three times, 3 min/time. The sections were blocked with 3% BSA at room temperature for 30 min. The sections were incubated with a rabbit anti-MTL antibody (1:1,000) and rabbit anti-MTLR (1:200) overnight at 4°C, followed by a secondary antibody conjugated with HRP and CY3-TSA (1:500, servicebio) or a goat anti-rabbit secondary antibody conjugated with 488 (1:400). In order to identify the expression of MTLR on neurons of the dorsal horn, double immunofluorescence staining for both MTLR and Tubb3 was performed simultaneously with rabbit anti-MTLR antibody (1:200) and mouse anti-Tubb3 antibody (1:100) overnight at 4°C. The sections were rinsed in PBS and incubated in goat anti-rabbit IgG coupled to if488 (1:400) and Goat anti-mouse IgG coupled to Cy5 (1:400) in the dark for 50 min at room temperature. The nuclei were counterstained with DAPI. After washing with PBS, all the sections were mounted on gelatin-coated slides and coverslipped with glycerol. Images were captured using a laser-scanning confocal microscope (TCS SP2; Leica, Wetzlar, Germany). Image-Pro Plus 6.0 (Media Cybernetics, Inc., Rockville, MD, USA) was used for analysis.

To measure locomotor function, we performed open field testing in which rats were placed in the center of a 100 × 100 cm chamber and locomotor activity was recorded by an overhead webcam connected to a laptop computer, and animals’ movements were automatically tracked for 30 min using Visualtrack. The total distance traveled and mean speed during the 30-min period were analyzed.

The data were expressed as mean ± SEM. Statistical analyses were performed using the unpaired Student’s t-test or one- or two-way ANOVA in behavioral experiments (ANOVA with repeated measures). All statistical analyses were performed using GraphPad Prism 9 (GraphPad Software Inc., San Diego, CA, USA). Pearson correlation analysis was performed between the MTL levels and MWT or TWL at each time point. Two-sided P-values < 0.05 were considered statistically significant.

There were no significant differences in MWT between the C and P groups at T0. Compared with the C group, the animals in the P group showed significantly decreased MWT at T1, T2, and T3, especially at T1. MWT gradually increased from T2 and returned close to baseline levels at T4 (Figure 1A). There were no significant differences in TWL between the C and P groups at T0. Compared with the C group, the P group showed significantly decreased TWL at T1, T2, and T3, especially at T1. TWL gradually increased from T2 and returned close to the baseline levels at T4 (Figure 1B). The results suggest that in the P group, the severest pain was observed at T1, which was alleviated at T4.

In order to verify whether MTL and MTLR are found in the spinal cord and what is their specific distribution, the changes in MTL and MTLR were examined by immunofluorescence at T0, T1, and T2 in the P group. MTL (red), MTLR (green), and neuronal markers were found in the superficial layer of the dorsal horn in the spinal cord. MTL and MTLR, as well as β-tubulin III and MTLR, were co-labeled in the superficial layers of the dorsal horn (Figure 2). The evidence suggests that MTL and MTLR interact in the cells located in the superficial layers of dorsal horn in the state of acute pain and provides a histological basis for possibilities that MTL and MTLR may play biological roles in the specific sensory regulation region. However, the current results only managed to indicate the existence of MTLR on dorsal horn neurons. The results also showed that with the decrease in pain threshold, the expression of MTLR in the superficial layers of the dorsal horn was upregulated. When the pain was most obvious at T1, the expression of MTLR was significantly higher than at T0, whereafter, the expression was downregulated at T2 but still higher than at T0 (Figure 3).

In order to observe the relation between the degree of pain and the central and peripheral gastric dynamic expression levels, we collected the stomach, duodenum, plasma, and spinal cord of rats in the C and P groups at T0-T4 to test MTL levels. There were no differences between groups C and P regarding the MTL levels in the stomach, duodenum, plasma, and spinal cord at T0. Compared with T0, the MTL levels in the gastric body of group P decreased at T1 and gradually increased at T2–T4 (Figure 4A). Compared with T0, the duodenal MTL levels were increased at T1 and gradually decreased at T2–T4 (Figure 4B). Compared with T0, the plasma MTL levels were decreased at T1, began to increase at T2, and gradually increased at T2–T4 (Figure 4C). In the spinal cord, compared with group C, MTL levels were increased at T1 and gradually decreased at T2–T4 (Figure 4D). Therefore, the expression of MTL varies in time after pain stimulation and varies according to organs/tissues. In the spinal cord, MTL varied with time after pain stimulation.

The expression of MTL was positively correlated with MWT and TWL in the gastric body (MWT: R² = 0.9087; TWL: R² = 0.8711), plasma (MWT: R² = 0.8255; TWL: R² = 0.8092). The expression of MTL in the duodenum (MWT: R² = 0.8646; TWL: R² = 0.8508) and spinal cord (MWT: R² = 0.8114; TWL: R² = 0.8075) were negatively correlated with MWT and TWL (Figure 5). Therefore, MTL expression is correlated with the changes of pain thresholds.

Compared with T0, the expression of MTL and MTLR in group P was increased significantly at T1 and T2, corresponding to the time points of ongoing acute pain. At T4, the behavioral pain indexes tended toward baseline levels, and the expression of MTL and MTLR gradually decreased and were not significantly different from baseline (Figure 6). Therefore, the expression of MTL and MTLR is a dynamic process in response to acute pain.

The behavioral results (Figure 7) showed that intrathecal injection of 1, 5, or 25 μg of MTL could increase the pain threshold of rats with incisional pain at T4, suggesting that MTL had a definite and long-lasting analgesic dose-dependent effect. It is unclear whether there is a ceiling effect to this dose-dependence for there were currently no experimental group with a higher dose of MTL.

The behavioral results (Figure 8) showed that the total distance traveled and mean velocity of rats with intrathecal injection of 25 μg of MTL did not differ significantly from the rats with the injection of normal saline during the 30-min period of the open field testing at 2 h after injection. The findings provide evidence that 25 μg of MTL intrathecal injection does not compromise locomotor function of rats.