Table of Contents Author Guidelines Submit a Manuscript
International Journal of Endocrinology
Volume 2013 (2013), Article ID 532850, 11 pages
Research Article

Calcium Activity of Upper Thoracic Dorsal Root Ganglion Neurons in Zucker Diabetic Fatty Rats

1Department of Drug Design and Pharmacology, Faculty of Health and Medical Sciences, University of Copenhagen, 2100 Copenhagen, Denmark
2Non-Clinical Development, Novo Nordisk A/S, 2760 Maaloev, Denmark

Received 14 December 2012; Revised 19 February 2013; Accepted 20 February 2013

Academic Editor: Gunnar Gislason

Copyright © 2013 Marie Louise Ghorbani et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


The aim of the present study was to examine the calcium activity of C8-T5 dorsal root ganglion (DRG) neurons from Zucker diabetic fatty rats. In total, 8 diabetic ZDF fatty animals and 8 age-matched control ZDF lean rats were employed in the study. C8-T5 dorsal root ganglia were isolated bilaterally from 14 to 18 weeks old rats, and a primary culture was prepared. Calcium activity was measured ratiometrically using the fluorescent Ca2+-indicator Fura-2 acetoxymethyl ester. All neurons were stimulated twice with 20 mM K+, followed by stimulation with either 0.3 or 0.5 μM Capsaicin, alone or in combination with algogenic chemicals (bradykinin, serotonin, prostaglandin E2 (all 10−5 M), and adenosine (10−3 M)) at pH 7.4 and 6.0. Neurons from diabetic animals exhibited an overall increased response to stimulation with 20 mM K+ compared to neurons from control. Stimulation with Capsaicin alone caused an augmented response in neurons from diabetic animals compared to control animals. When stimulated with a combination of Capsaicin and algogenic chemicals, no differences between the two groups of neurons were measured, neither at pH 7.4 nor 6.0. In conclusion, diabetes-induced alterations in calcium activity of the DRG neurons were found, potentially indicating altered neuronal responses during myocardial ischemia.

1. Introduction

It is estimated that 366 million people worldwide in the age of 20–79 years were affected by diabetes in 2010 and the prevalence is expected to increase to 552 million people in 2030 [1]. Cardiovascular disease is the main cause of death among people with diabetes, accounting for 50% of all diabetic deaths [1]. Diabetes mellitus is per se a risk factor for developing cardiovascular disease, with ischemic heart disease being the most frequently found [2], but also diabetic neuropathy, specifically cardiovascular autonomic neuropathy, is considered a risk factor [36]. Silent myocardial ischemia and infarction as well as sudden death are also more frequently found in patients where diabetes and cardiovascular autonomic neuropathy coexist and cardiovascular autonomic neuropathy is furthermore believed to be a contributing factor to the higher mortality and morbidity found in patients with diabetes mellitus following myocardial ischemia or infarction [37].

Neurons that can contribute to the pathological picture of autonomic neuropathy are the afferent sensory nerves coming from the organ, the neurons involved in central processing of the nociceptive signal, and the efferent autonomic nerves leading to the organ and regulating its function [8].

During myocardial ischemia, lactic acid is produced, causing a decrease in local pH (decrease of epicardial pH from 7.35 to 6.98 after 5 min of coronary artery occlusion has been demonstrated) [9], and release of inflammatory and/or algogenic (i.e., pain-causing) mediators, such as bradykinin, adenosine, serotonin, prostaglandin E2, and histamine [10]. These metabolic components have been shown to activate/sensitize spinal and vagal afferents in the heart. The effect is (1) release of neuropeptides leading to local effects in the myocardium and (2) transmission of the nociceptive signal through the primary afferents (via the dorsal root ganglia (DRGs)) to segments C8-T9 in the spinal cord [1115] or to the nucleus of the solitary tract in medulla oblongata. Neurons can project to the brain giving rise to perception of anginal pain and associated feelings (e.g., nausea and anxiety) and/or activate autonomic efferent fibers, leading to regulation of cardiovascular function [11].

Diabetes-induced alterations in cardiac sensory nerves could thus have pronounced effects on the way a patient responds to myocardial ischemia and infarction, including changes in perception of the pain often accompanying myocardial ischemia and infarction, and in the cardiac compensatory responses.

Diabetic neuropathy is thought to be caused by several hyperglycemia-induced changes: neurovascular dysfunction, lack of trophic support, and metabolic changes, such as an increased polyol pathway flux, increased production of advanced glycation end products (AGEs), and oxidative stress [16]. Furthermore, alterations in [Ca2+]i homeostasis have been suggested to contribute to the pathogenesis of diabetic neuropathy [1618]. Altered calcium channel activity has been shown in primary [18] and secondary sensory neurons from diabetic rodents [19].

Very little is known of the consequences of diabetes on sensory nerve function of primary afferents innervating the heart. Schultz found a decreased activity of cardiac vagal afferents in STZ-diabetic rats compared to control rats after epicardial application of bradykinin and capsaicin [20] and recently, diabetes-induced alterations in the spinal processing of nociceptive input from the heart were found in diabetic rats following cardiac stimulation [21].

The purpose of the present study was to investigate the calcium activity of C8-T5 dorsal root ganglion (DRG) neurons (primary neuron) in Zucker diabetic fatty rats, a spontaneous animal model for type 2 diabetes, and the effect of stimulation with algogenic substances, known to play part during myocardial ischemia, as well as with capsaicin, the pungent ingredient in chili peppers, at pH 7.4 and 6.0. To the best of our knowledge, this is the first study assessing the calcium activity in upper thoracic DRG neurons from any type of diabetic rat.

2. Methods and Materials

2.1. Animals

Animal procedures were approved by the Danish Animal Inspectorate (license no. 2004/561-943).

Experiments were performed in male diabetic Zucker diabetic fatty (ZDF) rats as well as age-matched nondiabetic ZDF rats. The ZDF rat is a spontaneous animal model for type 2 diabetes in which the animals are homozygous for nonfunctional leptin receptors [22, 23]. On a Purina 5008 diet, hyperglycemia will develop as early as an age of approximately 7 weeks [23].

The animals were stabled in pairs and were fed with Purina 5008, an energy-rich fodder. Once a week, the animals were weighed and blood glucose was measured from a tail vein using a glucometer (OneTouch Ultra2, Lifescan).

2.2. Primary Culture

At the age of 14–18 weeks, the rats were decapitated and C8-T5 DRG’s were removed bilaterally and placed in Hank’s Buffered Salt Solution (HBSS).

The ganglia were transferred to 5 mL papain solution (2 mg/mL, Sigma-Aldrich) for 35 min at 37°C. After centrifugation (2800 rpm, 2 min) the supernatant was removed and the pellet was resuspended in 5 mL of a collagenase/dispase solution (Collagenase: 1.11 mg/mL, Medinova; Dispase: 1.33 mg/mL, Roche Diagnostics GmbH). The solution was kept at 37°C for 20 min with routinely mixing. After centrifugation (2800 rpm, 2 min) and removal of supernatant, the cells were triturated in 1 mL HBSS (by at least 10 passages through a fire-polished Pasteur pipette) and gently placed on 7 mL percoll (20% v/v, Sigma-Aldrich). After centrifugation (2800 rpm, 7 min), the percoll containing ganglia remnants was removed and the pellet resuspended in 1 mL complete F-12 medium. After centrifugation (2800 rpm, 3 min), the supernatant was removed leaving approximately 30–60 μL of fluid. The cells were resuspended in the remaining fluid and approximately 10–20 μL of the cell suspension was placed on each of three poly-d-lysine/laminin 12 mm coverslips (Becton Dickinson CDC). The coverslips were placed at 37°C for two hours to allow the cells to adhere to the coverslips and afterwards, 2.5 mL of complete F-12 medium was added to each well. The cells were kept at 37°C until use (2–12 hours).

2.3. Calcium Imaging

The cells were transferred to PSS, adjusted to pH 7.4 and loaded with the fluorescent Ca2+-indicator Fura-2 acetoxymethyl ester (Fura-2/AM, 4.4 μM) dissolved in DMSO and Chremaphor EL detergent (4 : 1) for 30–60 min at 37°C. After loading, the cells were transferred to fresh PSS buffer and allowed to stand for approx. 30 min in order to ensure completion of the cytoplasmatic dye deesterification [24].

Changes in intracellular calcium concentrations in neurons from C8-T5 DRG’s were measured by FURA-2 ratiometric imaging in the acutely dissociated culture. At the time of measurement, cells were round and without processes.

The cells were illuminated alternately with light of 340 nm (specific for Ca2+-bound Fura-2) and 380 nm (specific for Ca2+-free Fura-2), and light of 510 nm was collected via a cooled CCD camera (Hamamatsu). The photomultiplier was coupled to a computer for data acquisition.

2.4. Experimental Protocols

Six different protocols were employed in the study, only one protocol per coverslip. Each coverslip was initially stimulated twice with 20 mM K+ for 10 s separated by 7 min wash with PSS. The first depolarization was undertaken to ensure vitality of the neurons and saturation of the intracellular compartments and was excluded from further analysis. After washout of the second administration of K+, neurons were stimulated for 10 s with one of the following solutions: (1) 0.3 μM capsaicin, (2) 0.5 μM capsaicin, (3) mixture of algogenic chemicals (AC) combined with 0.3 μM capsaicin, (4) AC combined with 0.5 μM capsaicin, (5) AC combined with 0.3 μM capsaicin at pH 6.0, or (6) AC combined with 0.5 μM capsaicin at pH 6.0. After stimulation with one of these solutions, the neurons were washed with PSS for 8 min. The experiment was concluded with a third K+ stimulation to evaluate the continued viability of the neuron for the extent of the protocol. The mixture of algogenic chemicals contained bradykinin, serotonin, prostaglandin E2, all in the concentration of 10−5 M, and adenosine (10−3 M). All substances used for stimulation were obtained from Sigma-Aldrich.

pH 6.0 was chosen for the two acidic protocols studies as DRG neurons containing TRPV1 and acid sensing ion channels (ASICs) have been demonstrated to react to this pH [2527].

2.5. Analysis

Prior to experiments, inclusion criteria for data analysis were defined: (1) labeling with Fura-2, (2) only neurons included, (3) neurons should stay within its marked area throughout the experiments, (4) cells responding to 20 mM K+ were included (≥20% increase from baseline), and (5) only neurons returning to baseline during washout period were included.

To monitor the changes in single cells, specific areas of interest were chosen for cells and for background. For each time frame, the ratio of background-corrected fluorescence intensities detected at 510 nm and resulting from excitation at 340 and 380 nm was calculated: This ratio is an arbitrary value and was used as a measure of the activity level and to record changes in the activity level of the neurons.

Baseline, peak value, and AUC of light intensity ratios were measured for each response in Chart 5 (AD Instruments). For K+ stimulations, AUC was calculated from 0 s to 180 s after initiation of response (); for all other stimulations, was calculated, where 0 s indicates beginning of rise in . The maximum increase from baseline, that is, the peak height, was calculated for all neurons, both in absolute terms (Δ), and relative as the percentage increase from baseline . For all stimulations except K+, Δ, and AUC were furthermore compared to the K+ response by calculating the ratio of a given response and the preceding K+ response for each neuron.

Neuron diameter was measured and the neurons were classified as small (S, <30 μm), medium (M, 30–39 μm), or large (L, ≥40 μm), similar to the classification by Lu et al. [28]. To further analyze the distribution of neuron diameter in control and diabetic animals, the neurons were divided into bins, each of 5 μm width.

Cells responding to 20 mM K+ were included in the analysis as neurons [18]. Responsiveness was defined as ≥ 20% increase from baseline (i.e., ≥ 20%), after stimulation with any substance in the protocol.

During analysis it was found that neurons stimulated with 0.3 and 0.5 μM Capsaicin elicited similar responses, and this was also seen for neurons stimulated with the combination of AC and either 0.3 or 0.5 μM Capsaicin. For subsequent analysis, 0.3 and 0.5 μM Capsaicin was therefore combined and henceforward is referred to as “Capsaicin” and furthermore, the protocols of AC in combination with either 0.3 or 0.5 μM Capsaicin were combined and henceforward are referred to “AC + Capsaicin.”

2.6. Statistical Analysis

The results are presented either as mean ± SEM or as median (5th percentile; 95th percentile), depending on normality of data. Comparisons were made with Student’s t-test or, when appropriate, the non-parametric Mann-Whitney Rank Sum Test. Comparisons of diameter distribution were done using Chi-square test. Statistical tests were performed with SigmaStat 3.11 and GraphPad Prism 5. was used as level of significance.

3. Results

3.1. Animals

In total, 8 diabetic ZDF animals and 8 age-matched control ZDF rats were used in this study. At the age of 7 weeks, the ZDF diabetic rats had developed significantly higher blood glucose concentrations than control rats and this hyperglycemia persisted for the whole experimental period. At the time of experiment, all diabetic animals had an elevated blood glucose level of  mM compared to  mM in control animals (, ). The body weight was larger in diabetic animals from 7 weeks of age and this increased body weight persisted throughout week 14 after which body weight no longer differed between control and diabetic animals. At the time of experiment, body weight was  g for the diabetic animals and  g for the control animals (, ).

3.2. Neurons

The mean cell diameter of the recorded DRG neurons was slightly smaller for diabetic than for control animals (25 μm (18.0; 33.2) versus 26 μm (19.0; 33.0), , ).

The distribution of small-, medium-, and large-sized neurons, though, did not differ between the two animal groups; for neurons from diabetic animals, 82% were small-sized, 17% medium-sized, and 1% large-sized compared to 83%, 15%, and 2%, respectively, for neurons from control animals. The distribution based on neuron diameter was also not significantly different in control and diabetic animals (Figure 1). When differentiating, however, between neurons responding to either of the capsaicin protocols (capsaicin-responsive) and neurons not responding to the tested capsaicin protocol (capsaicin-unresponsive), analysis showed that the distribution of “capsaicin-responsive” neurons from diabetic animals was significantly different from the distribution of control animals () with a median neuron diameter in diabetic animals of 23 μm (16.7; 30.0) compared to 25 μm (18.9; 34.1) in control animals ().

Figure 1: Distribution of neuronal cell diameter. The neurons were divided into 5 μm bins. Small-diameter neurons had diameters < 30 μm, medium-sized neurons had diameters between 30–39 μm, and large-diameter neurons had diameters of ≥ 40 μm. The majority of tested neurons were small-sized. n (control) = 552, n (diabetic) = 586.

The baseline Ca2+ level did not differ between diabetic and control animals (0.58 (0.46; 0.80) versus 0.59 (0.47; 0.77), , ). Response to a given stimulus was elicited by an increase in intracellular Ca2+-level and viewed as an increase in fluorescence intensity ratio, that is, a peak that slowly returned to baseline. An example of a capsaicin-responsive and-unresponsive neuron following stimulation with 20 mM K+ and a capsaicin-containing stimulus is depicted in Figure 2. Neuron responses with and without shoulder were found among both groups of animals as well as among capsaicin-responsive and -unresponsive neurons and thus, presence or lack of shoulder in the response was not confined to single groups of neurons investigated.

Figure 2: Example of calcium activity measurement. The figure depicts the trace of two cells obtained during a single experiment: a capsaicin-responsive (dark purple) and a capsaicin-unresponsive (light purple) neuron. The three small photomicrographs show the same neurons, at different stages of the experiment: (1) before stimulation, (2) during stimulation with 20 mM K+, and (3) during stimulation with 0.5 μM Capsaicin + algogenic chemicals. Black arrows beneath the trace indicate the beginning of stimulation. Time scale bar; 120 s. Cap; capsaicin.
3.3. K+-Induced Depolarization

Control as well as diabetic neurons responded to stimulation with 20 mM K+ by an increase in , assessed by an increase in 340/380 nm fluorescence ratio. The intracellular Ca2+ level returned to baseline during the washing period. A representative trace of the time- relationship is shown in Figure 3. The diabetic neurons showed an overall increased responsiveness to stimulation with 20 mM K+ compared to neurons from control animals (Table 1). This was seen as an augmented AUC, peak height (), and % increase from baseline . The increased responsiveness was observed both in “capsaicin-responsive” and “capsaicin-unresponsive” neurons (Table 2). For both control and diabetic animals it was seen that capsaicin-responsive neurons generally had a higher responsiveness to 20 mM K+ than capsaicin-unresponsive neurons (Table 2).

Table 1: Quantitative response of all neurons to 10 s stimulation with 20 mM K+. Results are shown as median (5%–95% percentile).
Table 2: Quantitative response to 10 s stimulation with 20 mM K+. The results were subcategorized according to whether they responded to the capsaicin protocol the respective neuron was stimulated with. Results are shown as median (5%–95% percentile).
Figure 3: Example of time- ratio development in dorsal root ganglion neurons from control and diabetic rats. The neurons were stimulated with 20 mM K+. For clarity in the illustration, the traces were background corrected. Arbitrary scale; vertical scale: 0.20 (340/380 nm light intensity ratio), and horizontal scale: 20 s.
3.4. Capsaicin Protocols

Stimulation with capsaicin alone, 0.3 or 0.5 μM (no combination) caused a rise in [Ca2+]i for approximately 23% of the neurons in both groups of animals. The neurons from diabetic animals exhibited an augmented response, assessed by a larger peak height, percentage increase from baseline, and AUC (Figures 4(a)4(c) and Table 3).

Table 3: Results from stimulation with capsaicin-containing solutions. Results are shown as Median (5th percentile; 95th percentile).
Figure 4: Traces and box plots from capsaicin experiments. Panels (a), (d), (g), and (j) show traces (examples) of the time- development for control and diabetic dorsal root ganglion neurons. The arrows indicate the onset of 10 sec stimulation with (a) capsaicin, (d) combination of Capsaicin and algogenic chemicals (AC); (g) combination of 0.3 μM Capsaicin and AC at pH 6.0, and (j) 0.5 μM Capsaicin and AC at pH 6.0. The traces were background corrected. As a measure of , the vertical axis is shown as the 340/380 nm light intensity ratio. (a) Vertical scale: 0.10; horizontal scale: 20 s, (d) vertical scale: 0.20; horizontal scale: 20 s.; (g) vertical scale: 0.10; horizontal scale: 20 s., and (j) vertical scale: 0.20; horizontal scale: 20 sec. Panels (b), (e), (h), and (k) display box plots on the absolute area under the curve (AUC) derived from stimulation with the in (a), (d), (g), and (j) mentioned capsaicin-containing solutions. Panels (c), (f), (i), and (l) display box plots with median, 25th and 75th percentiles on the relative AUC of the capsaicin-containing solution-derived increase and the preceding 20 mM K+-derived increase. The error bars indicate the 10th and 90th percentiles. Values below the 10th and above the 90th percentiles are shown as dots. + indicates the mean. *.

Capsaicin-responsive neurons stimulated with capsaicin in combination with the algogenic substances at either pH 7.4 or 6.0 were followed by increases in in neurons from both diabetic and control animals (Figures 4(d), 4(g), and 4(j)). None of the stimulations gave rise to differences in peak height or AUC between diabetic and control animals (Figures 4(d)4(k), Table 3).

Scatter plots of the capsaicin responses indicated that neurons responding to capsaicin-containing solutions might be divided in high- and low-responders (Figure 5).

Figure 5: Scatter plots displaying AUC for neurons from control and diabetic animals derived from stimulation with (a) capsaicin, (b) combination of Capsaicin and algogenic chemicals (AC); (c) combination of 0.3 μM Capsaicin and AC at pH 6.0, and (d) 0.5 μM Capsaicin and AC at pH 6.0.

Stimulation with 0.5 μM capsaicin in combination with the algogenic chemicals at pH 6.0 caused a significantly augmented Ca2+ response compared to any of the other stimulations. This was seen for neurons from diabetic animals (Δ; 1.63 (0.16; 1.87), AUC; 63.51 (2.87; 97.95), compared to every other stimulation of diabetic neurons, ) as well as from control animals (Δ; 1.7 (0.21; 2.37); AUC; 72.13 (1.58; 117.24), compared to every other stimulation of control neurons, ). The responses were similar in neurons from control and diabetic animals ().

4. Discussion

The major findings of the present study were the following: (1) baseline were unaltered in C8-T5 DRG neurons from ZDF diabetic compared to control animals, (2) K+-induced and (3) capsaicin-induced depolarization caused augmented currents in neurons from diabetic animals.

4.1. Baseline Ca2+-Levels

The current study showed no differences in baseline in C8-T5 DRG neurons from diabetic and control animals. This is in accordance with several other studies performed on DRG neurons from STZ-diabetic mice and rats, db/db diabetic mice and BB/W rats [18, 2932], and on dorsal horn neurons from lumbar L6–L7 spinal cord segments from STZ-diabetic rats [19, 33]. Shutov et al. [34], however, have found an increased baseline in L4–L6 DRG neurons from STZ-induced diabetic rats at the same time as an unaltered baseline in DH neurons, suggesting a differential effect by diabetes on primary and secondary neurons [34]. The segmental location of the neurons investigated also appear to be affected differently by diabetes as neurons with longer axons are more susceptible to neuropathic changes than neurons with shorter axons [8]. This is supported by reports of an increase in baseline [Ca2+]i in STZ-diabetic DRG neurons extracted from L4–L6 levels where the neurons have longer axons because they innervate the hind paws [18, 34], while Huang and colleagues, in the same study, found no baseline changes in C5-L3 DRG neurons [18], which agrees with the results from the present study on C8-T5 DRG neurons.

4.2. K+-Induced Depolarization

The present study found an increased depolarization-induced current in DRG neurons from ZDF diabetic animals compared to control animals. The increase comprised an absolute and a percentage increase in amplitude from baseline and an increased . Hall et al. also found an increased elevation of in thoracic DRG neurons (d. 20–40 μm) after stimulation with 50 mM KCl for 6 s in BB/W rats [32]. Most other studies with DRG and DH neurons have not detected differences in the amplitude of elevation between diabetic and nondiabetic rats and mice [19, 2931, 3335]. An exception, however, is Huang et al. who found a decreased amplitude of depolarization-induced current in small DRG neurons from L4–L6 in STZ-diabetic animals (14-week duration) [18]. The decrease in amplitude which contrasts the findings of the present study could be explained by the increase in baseline that Huang et al. found in the DRG neurons from diabetic animals [18], which might have limited the potential change in in these neurons. The discrepancy between studies might also be related to the duration of diabetes since Huang et al. investigated neurons from STZ-diabetic rats with longer duration (8–14 weeks) of diabetes than what was seen in other studies. The type of diabetes might also influence calcium homeostasis differently, since the present study and the study by Hall et al., both of which found increases in amplitude were performed in ZDF and BB/W rats [32], respectively compared to the other studies in which STZ-diabetic animals were employed [18, 19, 2931, 3335].

There is a general agreement on a prolonged recovery following depolarization-induced currents in neurons from diabetic rats and mice, measured as recovery time, residual (60 or 90 s after depolarization), or, as in the present study, [18, 19, 2931, 34, 35]. This suggests that the diabetes-induced effect on recovery of is somewhat fundamental as the studies are based on both type 1 and type 2 diabetes, different rodent species, neuron sizes, segmental orientations of the neurons investigated, and on different parts of the nervous system (DRG or DH). The level of in the DRG neurons is the sum of mechanisms leading to Ca2+-entry into the intracellular space and mechanisms leading to removal of Ca2+ from the intracellular space [36]. During stimulation, intracellular calcium level rises because calcium enters the cell via plasmalemmal voltage- and ligand-gated, and store-operated Ca2+-channels, and also because calcium is released from the endoplasmatic reticulum [36, 37]. Kinetics of the recovery phase is affected by release of Ca2+ ions from the mitochondria.

Voltage-dependent calcium currents through L- and N-channels are enhanced in dorsal horn neurons of STZ rats in vitro [33]. This is in accordance with the study by Hall et al. who found a decreased inhibitory effect of the opioid agonist Dyn A in thoracic DRGs from BB/W diabetic rats [32] and an impaired inhibitory G-protein function in thoracic and lumbar DRGs from BB/W and STZ rats, thus contributing to an increased calcium influx [38].

Several mechanisms thus seem to contribute to the increased K+-induced depolarization found in the present study, and since these changes are seen in different species, diabetes models, and at different DRG levels, the impact of diabetes on neuronal calcium metabolism seems to be universal.

4.3. Capsaicin Protocols

The present study showed an increased elevation of following stimulation with capsaicin (pooled results for 0.3 and 0.5 μM) and this agrees with studies by Hall et al. and Hong and Wiley who found an increased peak in thoracic DRG neurons (20–40 μm) from BB/W rats and larger and longer-lasting inward currents in DRG neurons from STZ-diabetic rats, respectively, following application of 1 μM Capsaicin [32].

The expression of TRPV1 has been shown to be decreased in the hearts of STZ-diabetic mice [39] although Strecker and colleagues reported unaltered capsaicin-induced CGRP release from human right atrium from subjects with diabetes [40].

The findings of increased Ca2+-response to K+ and capsaicin show that diabetes causes functional changes in these neurons. The augmented response measured in neurons from diabetic animals following stimulation with capsaicin suggests that the function or expression of TRPV1 might be altered by diabetes. Song et al. have found a decreased expression of membrane TVPR1 in heart [41] which is supported by Hong and Wiley who found decreased TRPV1 expression in small L1–L6 DRG neurons [42]. They showed, however, in the same study that functional TRPV1 was increased in neurons from diabetic animals and furthermore, the acid-evoked (pH 5.0) transient, mediated by TRPV1, was increased in these neurons [42] which is in accordance with the increased capsaicin response seen in the present study.

The function of the PKC-induced phosphorylation, and thus sensitization, of TRPV1 [43] has been shown to be increased in lumbar DRG neurons from STZ diabetic rats [42].

Glutamate receptors (GluR) colocalized with TRPV1 on small diameter DRG neurons have been demonstrated to contribute to the activity of the capsaicin-induced TRPV1 sensitization [44] and GluR could possibly contribute to the finding in the present study that capsaicin-responsive neurons elicited larger Ca2+ responses than neurons from capsaicin-unresponsive neurons (the difference was seen both in diabetic as well as control neurons).

Scatter plots of the responses to capsaicin-containing solutions indicated that the tested neurons might be either high- or low-responders. It cannot be ruled out that diabetes has a differentiated effect on these two potentially different subpopulations of neurons.

When 0.5 μM capsaicin was applied in combination with AC at acidic pH 6.0, the calcium response was larger (amplitude and AUC) for neurons, regardless of diabetic state, compared to all other applied solutions. This is in accordance with other studies showing that the capsaicin-induced neurons activity can be amplified by lowering pH and by adding inflammatory substances, such as bradykinin 5-HT, PGE2, and adenosine [10, 13, 26, 27]. There was no difference, however, between neurons from control and diabetic animals.

5. Conclusion

The findings of increased Ca2+-response to K+ and capsaicin show that diabetes causes functional changes in the investigated DRG neurons. The augmented response measured in neurons from diabetic animals following stimulation with capsaicin suggests that the function or expression of TRPV1 might be altered which is supported by other studies.

This is, to the best of our knowledge, the first study to investigate calcium activity in DRGs from ZDF rats and the study shows that the diabetes-induced effects on peripheral neuron function have common characteristics with other diabetic animal models. The ZDF rat thus seems to be a relevant model for studying diabetes-induced neuronal alterations in DRG neurons.

Diabetic neuropathy is a very complex condition with multiple factors playing part in the pathophysiology. The present study suggests that diabetes induces changes in DRG neuron function and this might be a contributing factor to the altered pain perception and the altered neuronal response by cardiac neurons seen in diabetic patients during myocardial ischemia and infarction.


This work was supported by The Danish Heart Association, no. 05-04-B120-A476-22223, The Novo Nordisk Foundation (30/11/2006), the Lundbeck Foundation ( 44/03), and Hørslev-Fonden (2003).


  1. International Diabetes Federation. Diabetes Atlas, 2011,
  2. J. S. Alpert, S. R. Chipkin, and N. Aronin, “Diabetes mellitus and silent myocardial ischemia,” Advances in Cardiology, vol. 37, pp. 297–303, 1990. View at Google Scholar · View at Scopus
  3. A. I. Vinik, T. Erbas, M. A. Pfeifer, E. L. Feldman, M. J. Stevens, and J. W. Russell, “Diabetic autonomic neuropathy,” in Ellenberg & Rifkin's Diabetes Mellitus, D. Porte, R. S. Sherwin, and A. Baron, Eds., chapter 46, pp. 789–804, McGraw-Hill, 6th edition, 2003. View at Google Scholar
  4. C. R. Kahn, G. C. Weir, G. L. King, A. M. Jacobson, A. C. Moses, and R. J. Smith, Eds., Joslin’s Diabetes Mellitus, Lippincott Williams & Wilkins, Philadelphia, Pa, USA, 14th edition, 2005.
  5. A. I. Vinik and D. Ziegler, “Diabetic cardiovascular autonomic neuropathy,” Circulation, vol. 115, no. 3, pp. 387–397, 2007. View at Publisher · View at Google Scholar · View at Scopus
  6. M. T. Johnstone and G. P. Kinzfogl, “Diabetes and heart disease,” in Diabetes and Cardiovascular Disease, M. T. Johnstone and A. Veves, Eds., Contemporary Cardiology, Series edited by C. P.Cannon, chapter 28, pp. 579–628, Humana Press, 2nd edition, 2005. View at Google Scholar
  7. A. I. Vinik, R. E. Maser, B. D. Mitchell, and R. Freeman, “Diabetic autonomic neuropathy,” Diabetes Care, vol. 26, no. 5, pp. 1553–1579, 2003. View at Publisher · View at Google Scholar · View at Scopus
  8. P. K. Thomas and D. R. Tomlinson, “Diabetic and hypoglycemic neuropathy,” in Peripheral Neuropathy, P. J. Dyck, P. K. Thomas, J. W. Griffin, P. A. Low, and J. F. Poduslo, Eds., chapter 64, pp. 1219–1250, Saunders, 1993. View at Google Scholar
  9. H. L. Pan, J. C. Longhurst, J. C. Eisenach, and S. R. Chen, “Role of protons in activation of cardiac sympathetic C-fibre afferents during ischaemia in cats,” Journal of Physiology, vol. 518, no. 3, pp. 857–866, 1999. View at Publisher · View at Google Scholar · View at Scopus
  10. S. T. Meller and G. F. Gebhart, “A critical review of the afferent pathways and the potential chemical mediators involved in cardiac pain,” Neuroscience, vol. 48, no. 3, pp. 501–524, 1992. View at Publisher · View at Google Scholar · View at Scopus
  11. R. D. Foreman, “Mechanisms of cardiac pain,” Annual Review of Physiology, vol. 61, pp. 143–167, 1999. View at Publisher · View at Google Scholar · View at Scopus
  12. D. C. Kuo, J. J. Oravitz, and W. C. DeGroat, “Tracing of afferent and efferent pathways in the left inferior cardiac nerve of the cat using retrograde and transganglionic transport of horseradish peroxidase,” Brain Research, vol. 321, no. 1, pp. 111–118, 1984. View at Publisher · View at Google Scholar · View at Scopus
  13. C. J. Benson, S. P. Eckert, and E. W. McCleskey, “Acid-evoked currents in cardiac sensory neurons: a possible mediator of myocardial ischemic sensation,” Circulation Research, vol. 84, no. 8, pp. 921–928, 1999. View at Google Scholar · View at Scopus
  14. F. Cervero and J. M. A. Laird, “Visceral pain,” The Lancet, vol. 353, no. 9170, pp. 2145–2148, 1999. View at Publisher · View at Google Scholar · View at Scopus
  15. F. Cervero, “Sensory innervation of the viscera: peripheral basis of visceral pain,” Physiological Reviews, vol. 74, no. 1, pp. 95–138, 1994. View at Google Scholar · View at Scopus
  16. G. Biessels and W. H. Gispen, “The calcium hypothesis of brain aging and neurodegenerative disorders: significance in diabetic neuropathy,” Life Sciences, vol. 59, no. 5-6, pp. 379–387, 1996. View at Publisher · View at Google Scholar · View at Scopus
  17. A. Verkhratsky and P. Fernyhough, “Mitochondrial malfunction and Ca2+ dyshomeostasis drive neuronal pathology in diabetes,” Cell Calcium, vol. 44, no. 1, pp. 112–122, 2008. View at Publisher · View at Google Scholar · View at Scopus
  18. T. J. Huang, N. M. Sayers, P. Fernyhough, and A. Verkhratsky, “Diabetes-induced alterations in calcium homeostasis in sensory neurones of streptozotocin-diabetic rats are restricted to lumbar ganglia and are prevented by neurotrophin-3,” Diabetologia, vol. 45, no. 4, pp. 560–570, 2002. View at Publisher · View at Google Scholar · View at Scopus
  19. N. V. Voitenko, E. P. Kostyuk, I. A. Kruglikov, and P. G. Kostyuk, “Changes in calcium signalling in dorsal horn neurons in rats with streptozotocin-induced diabetes,” Neuroscience, vol. 94, no. 3, pp. 887–890, 1999. View at Publisher · View at Google Scholar · View at Scopus
  20. H. D. Schultz, “Cardiac vagal chemosensory afferents: function in pathophysiological states,” Annals of the New York Academy of Sciences, vol. 940, pp. 59–73, 2001. View at Google Scholar · View at Scopus
  21. M. L. Ghorbani, C. Qin, M. Wu et al., “Characterization of upper thoracic spinal neurons receiving noxious cardiac and/or somatic inputs in diabetic rats,” Autonomic Neuroscience, vol. 165, no. 2, pp. 168–177, 2011. View at Google Scholar
  22. D. Chen and M. W. Wang, “Development and application of rodent models for type 2 diabetes,” Diabetes, Obesity and Metabolism, vol. 7, no. 4, pp. 307–317, 2005. View at Publisher · View at Google Scholar · View at Scopus
  23. G. J. Etgen and B. A. Oldham, “Profiling of Zucker diabetic fatty rats in their progression to the overt diabetic state,” Metabolism, vol. 49, no. 5, pp. 684–688, 2000. View at Google Scholar · View at Scopus
  24. R. M. Paredes, J. C. Etzler, L. T. Watts, W. Zheng, and J. D. Lechleiter, “Chemical calcium indicators,” Methods, vol. 46, no. 3, pp. 143–151, 2008. View at Publisher · View at Google Scholar · View at Scopus
  25. D. C. Immke and E. W. McCleskey, “Protons open acid-sensing ion channels by catalyzing relief of Ca2+ blockade,” Neuron, vol. 37, no. 1, pp. 75–84, 2003. View at Publisher · View at Google Scholar · View at Scopus
  26. M. Kress, S. Fetzer, P. W. Reeh, and L. Vyklicky, “Low pH facilitates capsaicin responses in isolated sensory neurons of the rat,” Neuroscience Letters, vol. 211, no. 1, pp. 5–8, 1996. View at Publisher · View at Google Scholar · View at Scopus
  27. M. Kress, P. W. Reeh, and L. Vyklicky, “An interaction of inflammatory mediators and protons in small diameter dorsal root ganglion neurons of the rat,” Neuroscience Letters, vol. 224, no. 1, pp. 37–40, 1997. View at Publisher · View at Google Scholar · View at Scopus
  28. S. G. Lu, X. Zhang, and M. S. Gold, “Intracellular calcium regulation among subpopulations of rat dorsal root ganglion neurons,” Journal of Physiology, vol. 577, no. 1, pp. 169–190, 2006. View at Publisher · View at Google Scholar · View at Scopus
  29. E. Kostyuk, N. Pronchuk, and A. Shmigol, “Calcium signal prolongation in sensory neurones of mice with experimental diabetes,” NeuroReport, vol. 6, no. 7, pp. 1010–1012, 1995. View at Google Scholar · View at Scopus
  30. E. Kostyuk, N. Voitenko, I. Kruglikov et al., “Diabetes-induced changes in calcium homeostasis and the effects of calcium channel blockers in rat and mice nociceptive neurons,” Diabetologia, vol. 44, no. 10, pp. 1302–1309, 2001. View at Publisher · View at Google Scholar · View at Scopus
  31. A. Shmigol and E. P. Kostyuk, “Mechanisms responsible for calcium signal formation in murine primary sensory neurons: their impairment by experimentally evoked diabetes,” Neurophysiology, vol. 27, no. 5, pp. 261–269, 1995. View at Google Scholar
  32. K. E. Hall, A. A. F. Sima, and J. W. Wiley, “Opiate-mediated inhibition of calcium signaling is decreased in dorsal root ganglion neurons from the diabetic BB/W rat,” The Journal of Clinical Investigation, vol. 97, no. 5, pp. 1165–1172, 1996. View at Google Scholar · View at Scopus
  33. N. V. Voitenko, I. A. Kruglikov, E. P. Kostyuk, and P. G. Kostyuk, “Effect of streptozotocin-induced diabetes on the activity of calcium channels in rat dorsal horn neurons,” Neuroscience, vol. 95, no. 2, pp. 519–524, 1999. View at Publisher · View at Google Scholar · View at Scopus
  34. L. Shutov, I. Kruglikov, O. Gryshchenko et al., “The effect of nimodipine on calcium homeostasis and pain sensitivity in diabetic rats,” Cellular and Molecular Neurobiology, vol. 26, no. 7-8, pp. 1541–1557, 2006. View at Publisher · View at Google Scholar · View at Scopus
  35. E. Kostyuk, N. Svichar, V. Shishkin, and P. Kostyuk, “Role of mitochondrial dysfunction in calcium signalling alterations in dorsal root ganglion neurons of mice with experimentally-induced diabetes,” Neuroscience, vol. 90, no. 2, pp. 535–541, 1999. View at Publisher · View at Google Scholar · View at Scopus
  36. M. J. Berridge, M. D. Bootman, and H. L. Roderick, “Calcium signalling: dynamics, homeostasis and remodelling,” Nature Reviews Molecular Cell Biology, vol. 4, no. 7, pp. 517–529, 2003. View at Publisher · View at Google Scholar · View at Scopus
  37. E. Carafoli, L. Santella, D. Branca, and M. Brini, “Generation, control, and processing of cellular calcium signals,” Critical Reviews in Biochemistry and Molecular Biology, vol. 36, no. 2, pp. 107–260, 2001. View at Google Scholar · View at Scopus
  38. K. E. Hall, J. Liu, A. A. F. Sima, and J. W. Wiley, “Impaired inhibitory G-protein function contributes to increased calcium currents in rats with diabetic neuropathy,” Journal of Neurophysiology, vol. 86, no. 2, pp. 760–770, 2001. View at Google Scholar · View at Scopus
  39. Z. Wei, L. Wang, J. Han et al., “Decreased expression of transient receptor potential vanilloid 1 impairs the postischemic recovery of diabetic mouse hearts,” Circulation Journal, vol. 73, no. 6, pp. 1127–1132, 2009. View at Publisher · View at Google Scholar · View at Scopus
  40. T. Strecker, A. Dieterle, P. W. Reeh, M. Weyand, and K. Messlinger, “Stimulated release of calcitonin gene-related peptide from the human right atrium in patients with and without diabetes mellitus,” Peptides, vol. 27, no. 12, pp. 3255–3260, 2006. View at Publisher · View at Google Scholar · View at Scopus
  41. J. X. Song, L. H. Wang, L. Yao, C. Xu, Z. H. Wei, and L. R. Zheng, “Impaired transient receptor potential vanilloid 1 in streptozotocin-induced diabetic hearts,” International Journal of Cardiology, vol. 134, no. 2, pp. 290–292, 2009. View at Publisher · View at Google Scholar · View at Scopus
  42. S. Hong and J. W. Wiley, “Early painful diabetic neuropathy is associated with differential changes in the expression and function of vanilloid receptor 1,” The Journal of Biological Chemistry, vol. 280, no. 1, pp. 618–627, 2005. View at Publisher · View at Google Scholar · View at Scopus
  43. S. Mandadi, P. Armati, and B. D. Roufogalis, “Real-time translocation adn function of PKCbII isoform in response to nociceptive signaling via the TRPV1 pain receptor,” Pharmaceuticals, vol. 4, pp. 1503–1517, 2011. View at Google Scholar
  44. Y.-H. Jin, M. Takemura, A. Furuyama, and N. Yonehara, “Peripheral glutamate receptors are required for hyperalgesia induced by capsaicin,” Pain Research and Treatment, vol. 2012, Article ID 915706, 8 pages, 2012. View at Publisher · View at Google Scholar