Introduction

Diabetic distal symmetric sensorimotor polyneuropathy affects ∼50% of patients with diabetes mellitus, and is a leading cause of foot amputation [1]. Evidence for the important role of the highly reactive oxidant peroxynitrite [2, 3] in peripheral diabetic neuropathy (PDN) is emerging from both experimental [47] and clinical [810] studies. Accumulation of nitrotyrosine (NT), a footprint of peroxynitrite injury, has been found in peripheral nerve, vasa nervorum, spinal cord and dorsal root ganglion (DRG) neurons in animal models of both type 1 (insulin-dependent) and type 2 (non-insulin-dependent) diabetes [37, 1113] and high glucose-exposed cultured human Schwann cells [14]. Recent experimental studies with peroxynitrite decomposition catalysts revealed an important contribution of peroxynitrite-induced injury (so called nitrosative stress) to diabetes-induced motor and sensory nerve conduction deficits, small sensory nerve fibre dysfunction and degeneration, and autonomic neuropathy [37, 15]. Clinical studies revealed increased plasma NT levels and their correlation with endothelial dysfunction and redistribution of sudomotor responses, an early sign of sympathetic nerve dysfunction, in type 1 diabetic patients [810]. Furthermore, plasma peroxynitrite generation assessed by the pholasin chemiluminescence test correlated with the diabetic neuropathy impairment score of the lower limbs [10].

Peroxynitrite is a product of superoxide anion radical reaction with nitric oxide. The latter can be produced by several nitric oxide synthase isoforms, i.e. endothelial nitric oxide synthase, inducible nitric oxide synthase (iNOS), and neuronal nitric oxide synthase. Whereas all three isoforms have been localised in the peripheral nervous system [14, 16, 17], their individual contributions to peroxynitrite-induced injury in tissue targets for PDN and development of nerve fibre dysfunction and degeneration remain unknown. Here, we provide the first evidence of the key role of iNOS in diabetes-induced nitrosative stress in the peripheral nerve, nerve conduction deficit, and small-fibre sensory neuropathy.

Methods

Reagents

Unless otherwise stated, all chemicals were of reagent-grade quality, and were purchased from Sigma Chemical Company, St Louis, MO, USA. Rabbit polyclonal anti-NT antibody was purchased from Upstate (Lake Placid, NY, USA) and mouse monoclonal anti-poly(ADP-ribose) (PAR) from Trevigen (Gaithersburg, MD, USA). Secondary Alexa Fluor 488 goat anti-rabbit and Alexa Fluor 488 goat anti-mouse antibodies as well as Prolong Gold Antifade Reagent were purchased from Invitrogen (Eugene, OR, USA). Avidin/Biotin Blocking Kit, M.O.M. Basic Kit, VECTASTAIN Elite ABC Kit (Standard*), DAB Substrate Kit, and 3,3′-diaminobenzidine (DAB) were obtained from Vector Laboratories (Burlingame, CA, USA). Rabbit polyclonal anti-protein gene product 9.5 (PGP 9.5) (ubiquitin C-terminal hydrolase) antibody was purchased from Chemicon International (Temecula, CA, USA). Other reagents for immunohistochemistry were purchased from Dako Laboratories (Santa Barbara, CA, USA).

Animals and limitations of the model of mice with iNos gene deficiency

The experiments were performed in accordance with regulations specified by the National Institutes of Health ‘Principles of Laboratory Animal Care, 1985 Revised Version’ and the Pennington Biomedical Research Center Protocol for Animal Studies. Several breeding pairs of B6.129P2-Nos2 tm1Lau/J (iNos −/−) mice, C57Bl6/J background, were obtained from Jackson Laboratories (Bar Harbor, ME, USA; stock number 002609). The use of iNos −/− mice for neurological studies is associated with some limitations because iNos gene (also known as Nos2) deficiency is known to result in impaired spinal cord regeneration, abnormal oligodendrocyte morphology and increased demyelination after neurotoxicant treatment (http://jaxmice.jax.org/strain/002609.html). Nevertheless, iNos −/−mice have normal motor and sensory nerve conduction velocities (MNCV and SNCV), normal thermal and mechanical algesia and tactile response thresholds, and, for this reason, have been considered suitable for assessment of the role of iNOS in PDN.

For the main experiment, a colony of iNos −/− mice was established at Pennington Biomedical Research Center. Mature male C57Bl6/J mice were purchased from Jackson Laboratories and served as controls. All the mice were fed standard mouse chow (PMI Nutrition International, Brentwood, MO, USA) and had access to water ad libitum. Diabetes was induced by a single injection of streptozotocin (STZ), 100 mg kg−1 day−1, i.p., to non-fasted animals. Blood samples for glucose measurements were taken from the tail vein 3 days after STZ injection and the day before the animals were killed. The mice with blood glucose ≥13.8 mmol/l were considered diabetic. The injected mice that had blood glucose concentrations in the non-diabetic range were given low-dose STZ injections (40 mg kg−1 day−1, i.p.) until they developed hyperglycaemia (typically, one to three additional injections). At the end of the study (duration of diabetes of 6 weeks), the physiological and behavioural tests were performed in the following order: tactile responses to flexible von Frey filaments (first day), tail-pressure Randall–Sellito test (second day), thermal algesia by tail-flick test (third day), thermal algesia by paw withdrawal test (fourth day), thermal algesia by hot plate test (fifth day), SNCV and MNCV (sixth day). Measurements of MNCV and SNCV were performed in mice anaesthetised with a mixture of ketamine and xylazine (45 mg/kg body weight and 15 mg/kg body weight, respectively, i.p.).

Anaesthesia, killing and tissue sampling

The animals were sedated by CO2, and immediately killed by cervical dislocation. Sciatic nerves, DRG and foot pads were fixed in 10% neutral buffered formalin solution containing 4% (wt/vol.) formaldehyde (Signa-Aldrich, Saint Louis, MO, USA) for assessment of NT and PAR by immunofluorescence histochemistry and intraepidermal nerve fibre density by conventional immunohistochemistry. PAR abundance is a measure of PAR polymerase (PARP) activity [18, 19].

Specific methods

Physiological tests

Sciatic MNCV and hindlimb digital SNCV were measured as we have described elsewhere [3, 20]. A TCAT-2 Temperature Controller with a RET-3 Temperature probe and an HL-1 Heat Lamp (Physitemp Instruments, Clifton, NJ, USA) was used to maintain body and hindlimb temperature at 37°C.

Behavioural tests

Tactile responses were evaluated by quantifying the withdrawal threshold of the hind paw in response to stimulation with flexible von Frey filaments as we have described [13]. Tail pressure thresholds were registered with a Paw/Tail Pressure Analgesia meter for the Randall–Selitto test (37215; Analgesy-Meter, UGO-Basile, Comerio VA, Italy). Pressure increasing at a linear rate of 10 g with the cut-off of 250 g to avoid tissue injury, was applied to the base of the tail. The applied tail pressure that evoked biting or licking behaviour was registered by an analgesia meter and expressed in grams. Three tests separated by at least 15 min were performed for each animal, and the mean value of these tests was calculated.

For thermal algesia, the paw withdrawal latency in response to the radiant heat (15% intensity, which produced a heating rate of ∼1.3°C/s, cut-off time 30 s) was determined as we have described [57, 12, 13] using an IITC model 336 TG combination tail-flick and paw algesia meter (IITC Life Science, Woodland Hills, CA, USA) with a floor temperature ∼32–33°C (manufacturer’s set up). For assessment of tail-flick response latencies, the device was set at 40% heating intensity (heating rate ∼2.5°C/s) with a cut-off at 10 s. In the hot plate test (IITC Model 39 Hot Plate Analgesia Meter; IITC Life Science) the unit had a plate preset temperature of 55°C. In all three tests, at least three readings per animal were taken at 15 min interval, and the average was calculated.

Immunohistochemical studies

All sections were processed by a single investigator and evaluated blindly. Low-power observations of skin sections stained for PGP 9.5 were made using a Zeiss Axioskop microscope (Carl Zeiss Imaging, Thornwood, NY, USA). Colour images were captured with a Zeiss Axiocam HRc CCD camera at 1,300 × 1,030 resolution (Carl Zeiss Imaging). Low-power images were generated with a ×40 acroplan objective using the automatic capturing feature of the Zeiss Axiovision software (Ver. 3.1.2.1). Low-power observations of sciatic nerve and DRG sections stained for NT and PAR were made using a Zeiss Axioplan 2 imaging microscope. Colour images were captured with a Photometric CoolSNAPHQ CCD camera (Photometrics Sales, Tucson, AZ, USA) at 1,392 × 1,040 resolution. Low-power images were generated with a ×40 acroplan objective using RS Image 1.9.2 software (Photometrics Sales).

NT immunoreactivities in the sciatic nerve and DRG were assessed by immunofluorescence histochemistry. In brief, sections were deparaffinised in xylene, rehydrated in decreasing concentrations of ethanol and washed in water. For immunofluorescence histochemistry, rabbit polyclonal anti-NT antibody was used in a working dilution of 1:400. Primary antibody was omitted in negative controls. Secondary Alexa Fluor 488 goat anti-rabbit antibody was applied in a working dilution of 1:200. Sections were mounted in Prolong Gold Antifade Reagent. The intensity of fluorescence was quantified using ImageJ 1.32 software (National Institutes of Health, Bethesda, MD, USA) and expressed as mean ± SEM for each experimental group.

PAR immunoreactivity was assessed as described [20] with minor modifications. In brief, sections of peripheral nerve and DRG were deparaffinised in xylene, rehydrated in decreasing concentrations of ethanol and washed in water. Non-specific binding was blocked with the mouse Ig blocking reagent supplied with the Vector M.O.M. Basic Immunodetection Kit. Then mouse monoclonal anti-PAR antibody was diluted 1:100 in 1% BSA in TRIS-buffered saline (TBS), and applied overnight at 4°C in the humidity chamber. Primary antibody was omitted in negative controls. Secondary Alexa Fluor 488 goat anti-mouse antibody was diluted 1:200 in TBS and applied for 2 h at room temperature. Sections were mounted in Prolong Gold Antifade Reagent. At least ten fields of each section were examined to select one representative image. Representative images were microphotographed. The number of nuclei with identifiable PAR fluorescence was calculated for each microphotograph and expressed as mean ± SEM for each experimental group. To evaluate neuronal PAR accumulation, a percentage of DRG neurons with weak, moderate and intense PAR immunofluorescence was calculated for each experimental group.

Intraepidermal nerve fibre density (INFD) was assessed as described [5, 6, 12, 13]. Three randomly chosen 5 μm sections from foodpad skin of each mouse were deparaffinised in xylene, hydrated in decreasing concentrations of ethanol, and washed in water. Non-specific binding was blocked by 10% goat serum containing 1% BSA in TBS (DAKO, Carpinteria, CA, USA) for 2 h, and the Avidin/Biotin Blocking kit, according to the manufacturer’s instructions. Then, rabbit polyclonal anti-PGP 9.5 antibody was applied in a 1:2,000 dilution. Secondary biotinylated goat anti-rabbit IgG (H+L) antibody was applied in a 1:400 dilution, and the staining performed with the VECTASTAIN Elite ABC Kit (Standard*). For visualisation of specific binding sites, the DAB Substrate Kit containing DAB was used. Sections were counterstained with Gill’s haematoxylin, dehydrated and mounted in Micromount mounting medium (Surgipath Medical, Richmond, IL, USA). Intraepidermal nerve fibre profiles were counted blindly by three independent investigators, under an Olympus BX-41 microscope (Leeds Precision Instruments, Minneapolis, MN, USA), and the average values were used. Microphotographs of stained sections were taken on an Axioscop 2 microscope (Zeiss) at ×4 magnification, and the length of epidermis was assessed with the ImagePro 3.0 program (Media Cybernetics, Bethesda, MD, USA). An average of 2.8 ± 0.3 mm of the sample length was investigated to calculate a number of nerve fibre profiles per mm of epidermis.

Statistical analysis

Results are expressed as means ± SEM. Data were subjected to equality of variance F test, and then to log10 transformation, if necessary, before one-way ANOVA. Where overall significance (p < 0.05) was attained, individual between-group comparisons were made using the Student–Newman–Keuls multiple-range test. Significance was defined at p ≤ 0.05. When between-group variance differences could not be normalised by log transformation (datasets for body weights and plasma glucose), the data were analysed by the non-parametric Kruskal–Wallis one-way ANOVA, followed by the Bonferroni–Dunn test for multiple comparisons.

Results

Weight gain during the 6 week study was comparable in non-diabetic wild-type (17%) and non-diabetic iNos −/− (20%) mice (Table 1). Whereas diabetic wild-type mice lost 14% of their initial body weight, diabetic iNos −/− mice did not display any weight loss, and moreover, gained 5% of weight. Initial (after STZ injection) blood glucose concentrations were 72% and 79% higher in diabetic wild-type and diabetic iNos −/− mice compared with the corresponding controls. Hyperglycaemia progressed with the prolongation of diabetes, and the difference between final blood glucose concentrations in both groups and corresponding controls exceeded threefold. Final blood glucose concentrations were similar in diabetic wild-type and diabetic iNos −/− mice.

Table 1 Initial and final body weights and blood glucose concentrations in experimental groups

Sciatic MNCV and hindlimb SNCV were 20 and 14% lower in diabetic wild-type mice compared with non-diabetic controls (p < 0.01 for both comparisons, Fig. 1a,b). In contrast, diabetic iNos −/− mice preserved normal MNCV and SNCV.

Fig. 1
figure 1

Sciatic MNCV (a) and hindlimb digital sensory nerve conduction velocities (b) in control (C) and diabetic (D) wild-type and iNos −/− mice. Means ± SEM, n = 6–8 per group. **p < 0.01 vs corresponding non-diabetic groups; †† p < 0.01 vs diabetic wild-type mice

The latency of hind-paw withdrawal in response to radiant heat was increased by 103% in diabetic wild-type mice compared with the control group (p < 0.01), consistent with clearly manifest thermal hypoalgesia (Fig. 2a). This is in agreement with the results of tail-flick and hot plate tests, which also revealed increased thermal response latencies in the diabetic wild-type mice (Fig. 2b,c). In contrast, only very minor hypoalgesia (a 10% increase in the response latency) was registered with the hind-paw withdrawal test in diabetic iNos −/− mice. Note, however, that tail-flick response latencies were similarly increased in diabetic wild-type (21%) and diabetic iNos −/− (17%) mice compared with the corresponding non-diabetic controls. The hot plate test results were in the normal range in diabetic iNos −/− mice.

Fig. 2
figure 2

Paw withdrawal latencies in response to radiant heat (a), tail-flick test response latencies (b) and hot-plate test response latencies (c) in control (C) and diabetic (D) wild-type and iNos −/− mice. Means ± SEM, n = 8–11 per group. *p < 0.05 and **p < 0.01 vs corresponding non-diabetic groups; †† p < 0.01 vs diabetic wild-type mice

Diabetic wild-type and iNos −/− mice displayed moderate mechanical hypoalgesia detected with the tail pressure Randall–Sellito test (Fig. 3a). The tail pressure threshold was increased by 19% in diabetic wild-type mice and by 13% in diabetic iNos −/− mice compared with the corresponding non-diabetic groups (p < 0.05 and p > 0.05, respectively). The severity of tactile allodynia was lower in diabetic wild-type compared with diabetic iNos −/− mice (Fig. 3b). Tactile response thresholds were reduced by 70% in diabetic wild-type mice and by 40% in diabetic iNos −/− mice compared with the corresponding untreated groups (p < 0.01 for both comparisons).

Fig. 3
figure 3

Mechanical withdrawal thresholds in tail-pressure Randall–Sellito tests (a) and tactile response thresholds in response to stimulation with flexible von Frey filaments (b) in control (C) and diabetic (D) wild-type and iNos −/− mice. Means ± SEM, n = 6–11 per group. **p < 0.01 vs non-diabetic control mice; †† p < 0.01 vs diabetic wild-type mice

INFD was reduced by 36% in diabetic wild-type mice and by 14% in diabetic iNos −/− mice compared with the corresponding controls (p < 0.05 and p > 0.05, respectively, Fig. 4a,b). Note, however, that iNos gene deficiency was associated with a ∼10% decrease in INFD in non-diabetic mice.

Fig. 4
figure 4

Intraepidermal nerve fibre profiles in control (C) and diabetic (D) wild-type and iNos −/− mice. a Representative images of intraepidermal nerve fibre profiles, magnification ×200. Arrows indicate intra-epidermal nerve fibres. b Skin fibre density of control and diabetic wild-type and iNos −/− mice. Means ± SEM, n = 8–11 per group. *p < 0.05 vs control mice

NT immunofluorescence was increased by 69% in the sciatic nerves of diabetic wild-type mice compared with non-diabetic controls (p < 0.01, Fig. 5a,b). In contrast, no diabetes-induced NT accumulation was detected in the sciatic nerves of iNos −/− mice. NT immunofluorescence of DRG was increased by 56% in diabetic wild-type mice and by 39% in diabetic iNos −/− mice compared with the corresponding non-diabetic groups (p < 0.01 for both comparisons, Fig. 5c,d).

Fig. 5
figure 5

Representative microphotographs of immunofluorescent staining of NT in sciatic nerves (a) and DRG (c) of control (C) and diabetic (D) wild-type and iNos −/− mice. Magnification ×40. NT fluorescence counts in sciatic nerves (b) and DRG (d) of control and diabetic wild-type and iNos −/− mice. Means ± SEM, n = 7–11 per group. **p < 0.01 vs non-diabetic control mice; †† p < 0.01 vs diabetic wild-type mice. RFU, relative fluorescence units

The number of PAR-positive nuclei was 71% higher in the sciatic nerve of diabetic wild-type mice, compared with non-diabetic controls (Fig. 6a,b). The percentage of DRG neurons with weak PAR immunofluorescence was lower, and of those with moderate and intense immunofluorescence higher in diabetic wild-type mice compared with the corresponding control group (Fig. 6c,d). iNos gene deficiency reduced the percentage of neurons with moderate and intense PAR fluorescence and increased the percentage of neurons with weak PAR fluorescence in diabetic mice.

Fig. 6
figure 6

Representative microphotographs of immunofluorescent staining of PAR in sciatic nerve (a) and DRG neurons (c) of control (C) and diabetic (D) wild-type and iNos −/− mice. Magnification ×40. (b) The number of PAR-positive nuclei in sciatic nerve; d percentage of DRG neurons with weak, moderate, and intense PAR immunofluorescence. The number of DRG neurons with weak, moderate and intense PAR immunofluorescence was expressed as a percentage of neurons with identifiable PAR immunofluorescence (examples are shown by white arrows) in the dorsal root ganglia of control and diabetic wild-type and iNos −/− mice. Means ± SEM, n = 8–11 per group. **p < 0.01 vs non-diabetic control mice; †† p < 0.01 vs diabetic wild-type mice

Discussion

Evidence for the important role of iNOS in diabetic complications is emerging. Diabetes-associated iNOS upregulation has been found in the retina [21], heart [22], vascular endothelium [23] and smooth muscle layer [24]. Furthermore, high glucose-induced iNOS overexpression has been reported for human Schwann cells [14], human retinal and coronary artery endothelial cells [25, 26], and rat and murine glomerular mesangial cells [27, 28]. Advanced glycation end-products induced iNos expression via a p38 mitogen-activated protein kinase-dependent pathway [29]. PARP activation, another important player in the regulation of iNos gene expression, upregulated iNOS via nuclear factor-kappaB activation [30]. Studies in iNos −/− mice revealed an important role for iNOS in subnormal retinal oxygenation, leucostasis, blood–renal barrier breakdown and formation of pericyte ghosts and acellular capillaries characteristic of peripheral diabetic retinopathy [3133]. They have also implicated iNOS in diabetes-induced endothelial dysfunction [23], impaired vascular reactivity [24], cardiomyopathy [34], myocardial ischaemia–reperfusion injury [35] and stroke [36], as well as glomerulosclerosis and tubulointerstitial fibrosis characteristic of chronic diabetic nephropathy [37].

The role of iNOS in diabetic neuropathy has not been studied. iNos gene expression has been identified in rat peripheral nerves and dorsal root ganglia [16]. iNos mRNA expression was found to be reduced rather than increased in the sciatic nerve of rat models of both short-term and long-term diabetes [16]. The latter is consistent with the demonstration of a diabetes-associated decrease in iNos mRNA expression in penile intracavernous nerves [17]. Nevertheless, the present study unequivocally demonstrates a key role of iNOS in peroxynitrite injury to peripheral nerve, motor and sensory nerve conduction deficits, and small-fibre sensory neuropathy.

Our findings are consistent with other reports suggesting that nitrosative stress is a characteristic feature of experimental PDN [3840]. Accumulation of NT is clearly manifest in peripheral nerve and DRG neurons of wild-type STZ-diabetic mice, consistent with previous observations of our group and others made in STZ-diabetic rodents [4, 6, 7], as well as in ob/ob [12] and high-fat-diet fed [13] mice. Using specific markers for certain cell types of PNS, we localised immunoreactive NT in endothelial and Schwann cells of peripheral nerve, as well as neuronal and glial cells of dorsal root ganglia (V. R. Drel and I. G. Obrosova, unpublished results). Thus, nitrosative stress affects all major tissue targets for PDN. Diabetic iNos −/− mice did not display NT accumulation in peripheral nerve, but were not protected from nitrosative stress in DRG. Despite the latter circumstance, iNos −/− mice preserved normal MNCV and SNCV. Furthermore, iNos gene deficiency alleviated the severity of small-fibre sensory neuropathy. These findings suggest that iNOS-dependent peroxynitrite formation in axons and Schwann cells, rather than cell bodies, of peripheral nerve plays a major role in functional and structural changes of diabetic neuropathy.

Evidence for the important role of PARP activation, another phenomenon closely linked to oxidative–nitrosative stress, in diabetic complications is emerging [18, 41, 42]. We [20, 4345] and others [15] have demonstrated a key role of PARP activation in motor and sensory nerve conduction and nerve blood flow deficits, thermal hyper-and hypoalgesia, mechanical hyperalgesia, tactile allodynia, exaggerated flinching behaviour in the formalin pain test, and small sensory nerve fibre degeneration associated with PDN, as well as in diabetic autonomic neuropathy. PARP activation manifest by PAR accumulation was present in both sciatic nerve and DRG in diabetic wild-type mice. iNos gene deficiency completely prevented diabetes-induced peripheral nerve PAR accumulation, consistent with the lack of enhanced nitrosative stress. Of interest, despite the presence of peroxynitrite injury in DRG of diabetic iNos −/− mice, PAR accumulation in DRG neurons was markedly alleviated. The significance and mechanisms of this effect cannot be interpreted based on current knowledge. However, growing evidence suggests that PARP activation is not a mere consequence of oxidative–nitrosative stress, but can be mediated via other biochemical mechanisms, e.g. phosphorylation by extracellular signal regulated kinase [46].

In conclusion, iNOS plays a key role in peroxynitrite injury to peripheral nerve, MNCV and SNCV deficits and small-fibre sensory neuropathy. Nitrosative stress in axons and Schwann cells, rather than DRG neurons, plays a major role in peripheral nerve dysfunction and degeneration associated with PDN. The findings support the rationale for development of specific inhibitors of iNos for prevention and treatment of this devastating complication of diabetes mellitus.