Nitric oxide (NO) is a molecular diffusible messenger that plays different roles in the nervous system. NO is produced from l-arginine by the nitric oxide synthase (NOS), an enzyme which requires several cofactors including calcium, calmodulin, and nicotinamide adenine dinucleotide phosphate (NADPH). Three NOS isoforms have been identified and distinguished by their tissue distribution: neuronal (nNOS), inducible (iNOS), and endothelial (eNOS). The nNOS (Type I) is located in a discrete neuronal populations of the central (CNS) and peripheral (PNS) nervous system; the iNOS (Type II) is expressed by different cell types in response to inflammatory stimuli; the eNOS (Type III) has been identified in endothelial cells and astrocytes (Garthwaite, 1991; Bredt and Snyder, 1992; Dawson and Dawson, 1996; Moncada et al., 1997; Bredt, 1999; Davis et al., 2001; Ahern et al., 2002; Calabrese et al., 2007; Bryan et al., 2009). Interestingly, the nNOS is also present in macrophages and endothelial cells, in which it is probably involved in the basal modulation of vascular tone (Seddon et al., 2008). The NO-synthesizing neurons, or nitrergic neurons, can be revealed using both histochemical staining for NADPH-diaphorase (NADPH-d) and immunohistochemical detection of nNOS. The diaphorase activity and consequently its distribution in the CNS is superimposed to nNOS immunoreactivity (Dawson et al., 1991; Vincent and Kimura, 1992; Giraldi-Guimarães et al., 1999; Bombardi et al., 2011, 2006). NADPH-d and nNOS containing neurons have been localized in the spinal cord in several mammalian species, including mouse, rat, rabbit, dog, cat, sheep, monkey, and human (Brüning, 1992; Valtschanoff et al., 1992a, 1992b; Dun et al., 1992, 1993; Spike et al., 1993; Terenghi et al., 1993; Saito et al, 1994; Vizzard et al., 1994; Wetts and Vaughn, 1994; Pullen and Humphreys, 1995; López-Figueroa et al., 1996, 1997; Marsăla et al., 1998, 1999; Kluchová et al., 2000). Despite the previous studies, no information is available on the distribution of nitrergic neurons in the spinal cord of the cetaceans. Nitrergic innervation is important in the nervous regulation of blood vessels, since NO is a potent vasodilator (Gaw et al., 1991). Considering that the spinal cord of dolphins is surrounded by a dense mesh of blood vessels that constitute the huge spinal rete mirabilis (a feature characteristic of cetaceans), the nervous control of regional peri-spinal blood flow is of paramount importance. To further address this question, the present study determines the distribution and morphology of nitrergic spinal neurons in the bottlenose dolphin (Tursiops truncatus) using histochemical and immunohistochemical procedures.
Nitric oxide (NO) is a freely diffusible gaseous neurotransmitter generated by a selected population of neurons and acts as a paracrine molecule in the nervous system. NO is synthesized from l-arginine by means of the neuronal nitric oxide synthase (nNOS), an enzyme requiring nicotine adenine dinucleotide phosphate (NADPH) as cofactor. In this study, we used histochemical and immunohistochemical techniques to investigate the distribution of NADPH-diaphorase (NADPH-d) and nNOS in the spinal cord of the bottlenose dolphin (Tursiops truncatus). Cells with a fusiform-shaped somata were numerous in the laminae I and II. The intermediolateral horn showed darkly-stained cells with a multipolar morphology. Neurons with a multipolar or fusiform morphology were observed in the ventral horn. Multipolar and fusiform neurons were the most common cell types in lamina X. Nitrergic fibers were numerous especially in the dorsal and intermediolateral horns. The presence of nitrergic cells and fibers in different laminae of the spinal cord suggests that NO may be involved in spinal sensory and visceral circuitries, and potentially contribute to the regulation of the complex retia mirabilia. Anat Rec, 296:1603–1614, 2013. © 2013 Wiley Periodicals, Inc.
MATERIALS AND METHODS
Spinal cords were obtained from three specimens of bottlenose dolphins Tursiops truncatus from the collections of the Mediterranean marine mammal tissue bank of the University of Padova (Italy) (website: http://www.mammiferimarini.sperivet.unipd.it/eng/index.htm; CITES IT 020). The Bank, recognized by the Italian Ministry of the Environment, harvests tissues from cetaceans stranded along the Italian coastline or dead in captivity. Tissue samples are removed; coded for freshness; and either fixed in formalin or frozen. The spinal cords used in this study were dissected out within 24 hr after death, and divided into cervical (C), thoracic (T), lumbar (L), and caudal (Ca) parts. Each part was divided in the different spinal segments (C1–C8; T1–T12; L1–L16; Ca1–Ca25), and immediately fixed by immersion in cold 4% paraformaldehyde in 0.1 M phosphate buffer saline (PBS; pH 7.4), for at least 24 hr. After rinsing in PBS, the tissue was cryoprotected in a 30% sucrose solution in PBS (pH 7.4) at 4°C. The spinal segments were cut on the transverse, sagittal, and dorsal plane into sections having a thickness of 30 μm using a freezing sliding microtome. The sections were stored in a tissue collecting solution (30% sucrose in PBS containing 0.1% sodium azide) until processed, at 4°C for immunohistochemical and histochemical staining, or in 10% formalin at room temperature for thionin staining.
The free-floating transverse (five for each spinal segment), sagittal (five for each spinal segment), and dorsal (five for each spinal segment) sections were washed in PBS (three times for 10 min each). To eliminate endogenous peroxidase activity, the sections were treated with 1% H2O2 in PBS for 30 min at room temperature. Sections were rinsed in PBS (three times for 10 min each) and incubated in a solution containing 10% normal goat serum (Colorado Serum, Denver, CO, #CS 0922) and 0.5% Triton X-100 (Merck, Darmstadt) in PBS for 2 hr at room temperature to block nonspecific binding. Thereafter, the sections were incubated in a solution containing mouse monoclonal antibody anti-nNOS (1:500; sc-5302; Santa Cruz Biotechnology, CA) for 48 hr at 4°C. The primary antibody was diluted in 0.01 M PBS with the addition of 0.1% sodium azide, containing 1% normal goat serum and 0.5% Triton X-100. After washing in PBS (three times for 10 min each), the sections were incubated in goat biotinylated anti-mouse (1:200; BA-9200, Vector Laboratories, Burlingame, CA) for 2 hr at room temperature. The secondary antibody was diluted in PBS containing 1% normal goat serum and 0.5% Triton X-100. The sections were then transferred to avidin–biotin complex (ABC kit Vectastain, PK-6100, Vector Laboratories, Burlingame, CA) for 45 min, and the immunoperoxidase reaction was developed by 3,3′-diaminobenzidine (DAB kit, SK-4100, Vector Laboratories, Burlingame, CA). After washing, the sections were mounted onto gelatin-coated slides, dried overnight at 37°C, dehydrated in ethanol, cleared in xylene, and coverslipped with Entellan (Merck, Darmstaldt, Germany).
NADPH-Diaphorase Histochemical Reaction
This procedure was carried out on free-floating sections. Fifteen sections (five transverse, five sagittal, and five dorsal) for each spinal segment were incubated in 0.1 M Tris buffer (pH 8.0) containing 1 mM β-NADPH (Sigma N-1630), 0.1 mM nitroblue tetrazolium (Sigma N-6876), and 0.3% Triton X-100 for 1 hr at 37°C. After washing in 0.3% Triton X-100 in Tris-buffered saline, the sections were mounted onto coated slides, dried overnight, dehydrated in ethanol, cleared in xylene, and coverslipped with Entellan (Merck, Darmstaldt, Germany).
Procedures of Staining Control
The specificity of monoclonal antibody used in this study has been described previously (Bombardi et al., 2011). Incubation without the substrate NADPH or without the electron acceptor Nitroblue tetrazolium chloride (NBT) drastically reduced the intensity of NADPH-d staining. This aspect demonstrated the specificity of NADPH-d histochemical staining.
To help identify the laminae of the spinal cord, sections adjacent to immunoperoxidase and NADPH-diaphorase histochemical preparations were stained with thionin as follows. Sections were taken out of the 10% formaldehyde solution, mounted on gelatin-coated slides, and dried overnight at 37°C. Sections were defatted 1 hr in a mixture of chloroform/ethanol 100% (1:1), and then rehydrated through a graded series of ethanol, 2× 2 min in 100% ethanol, 2 min in 96% ethanol, 2 min in 70% ethanol, 2 min in 50% ethanol, 2 min in dH2O, and stained 30 sec in a 0.125% thionin (Fisher Scientific) solution, dehydrated and coverslipped with DPX (BDH Laboratory Supplies Poole, England).
Analysis of Sections
Sections stained using thionin, histochemical, and immunohistochemical methods were analyzed with a Leica DMRB microscope. The distribution of nitrergic somata in a representative series of transverse sections was plotted by means of a computer-aided digitizing system (AccuStage 5.1, St. Shoreview, MN). Camera lucida drawings from the adjacent thionin-stained sections were used to define the laminar and regional boundaries of the spinal cord. The outlines were superimposed on computer generated plots using Corel Draw X3 (Corel Corporation, Ottawa, Ontario, Canada). KS 300 Zeiss software (Kontron Elektronik, Germany) was utilized for morphometric analysis of the nitrergic neurons located in the cervical (C1–C8), thoracic (T1–T12), lumbar (L1–L16), and caudal (Ca1–Ca25) spinal segments. Only the neurons with an evident nucleus were included in the perikaryal area analysis. In each animal, the perikaryal areas, expressed as means ± SD, were measured in representative transverse, sagittal and dorsal sections of each spinal segment after manual tracing of their outline. In particular, for each section plane, we used two nonconsecutive sections obtained from each spinal segment. The contrast and brightness of the figures were adjusted to reflect the appearance of the labeling seen through the microscope using Adobe Photoshop CS3 Extended 10.0 software (Adobe Systems, San Jose, CA).
Using thionin staining we observed that the gray matter of the bottlenose dolphin spinal cord could be subdivided bilaterally in dorsal, lateral (intermediolateral cell column), and ventral horns (Fig. 1A). Our data confirm previous investigations reporting that the ventral horns were more developed than the dorsal ones throughout the entire craniocaudal extension of the spinal cord (see Flanigan, 1966; Morgane and Jacobs, 1972). Using the Rexed's method (Rexed, 1954) we could recognize ten laminae (layers), the gray matter of the spinal cord:
- Lamina I was thin and covered the upper surface of the dorsal horn. In transverse sections, this lamina showed especially small spheroidal neurons, which appeared elongated and spindle-shaped cells in sagittal and dorsal sections (Fig. 1B).
- Lamina II was located immediately ventral to the Lamina I and contained tightly packed small spheroidal (transverse sections) or fusiform (sagittal and dorsal sections) neurons (Fig. 1B).
- Lamina III was thicker than Lamina II and, as Lamina IV, contained scattered neurons of different sizes. The majority of the somata had a polygonal appearance (Fig. 1C).
- Lamina IV contained small spheroidal and polygonal somata varying from small to large.
- Lamina V had small, medium, and large neurons. These cells were not tightly packed and showed a polygonal morphology.
- Lamina VI was quite thin and contained fusiform and polygonal somata of different sizes.
- Lamina VII was well developed and, laterally, contained the intermediolateral cell column. This nucleus, which was extended from T1 to L4/L5, contained polygonal medium-sized somata (Fig. 1D).
- Lamina VIII was thin and showed small, medium, and large polygonal neurons (Fig. 1E).
- Lamina IX could be subdivided in two principal neuronal groups (lateral and ventral) where it was possible to identify medium and large polygonal somata corresponding to motoneurons (Fig. 1F).
- Lamina X, located in the central gray matter, exhibited spheroidal and polygonal from small to medium-sized neurons (Fig. 1G).
Since the limb enlargements were absent in the spinal cord of the bottlenose dolphin, the thickness of the different laminae was quite constant along the craniocaudal extension of the spinal cord.
NADPH-Diaphorase Activity and Neuronal Nitric Oxide Synthase Immunoreactivity in the Spinal Cord
The distribution of NADPH-d-positive cells in the spinal cord of the bottlenose dolphin appeared identical to that obtained for nNOS immunohistochemistry; however, only the NADPH-d-positive neurons had a typical Golgi-like appearance. Usually, the intensity of the staining was higher in histochemical than in immunohistochemical preparations. Prominent laminar differences were noted in the distribution of nitrergic neurons throughout the length of the spinal cord (Fig. 2). In Laminae I, II, VII, and X, many neurons were labeled (Fig. 2). The morphology and the size of the nitrergic neurons varied depending on their location within the gray matter. The population of nitrergic neurons could be divided into three major cell types based primarily on the morphology of the soma: polygonal, fusiform, and spheroidal. Polygonal cells were multipolar with angular soma from which arose at least three primary dendrites that varied in thickness. The somata of fusiform neurons had an ovoidal shape with primary dendrites emanating from the opposite poles of the soma. Since most fusiform cells exhibited only two primary dendrites (especially in Laminae I and II), these cells could be defined as bipolar neurons. Spheroidal cells had a round cell body and did not show any visible dendrites. The neuropil staining consisted of dendrites, axons, puncta, and diffuse staining.
Dorsal Horn (Laminae I–VI)
A large population of nitrergic neurons was found in the dorsal horn in all spinal segments. Labeled neurons were prevalently located in Laminae I and II (Fig. 3A–H). Most of these neurons were bipolar or multipolar, in sagittal and dorsal sections, and multipolar, bipolar, or spheroidal, in transverse sections (Fig. 3E–H). The somata of bipolar neurons had a fusiform shape with two primary dendrites emanating from the opposite poles of the soma (Fig. 3E). In sagittal and dorsal sections it was possible to appreciate that the principal axis of the fusiform neurons was directed rostro-caudally. The mean cross-sectional area of neurons located in Laminae I and II is reported in Table 1. Usually fusiform cells were smaller than multipolar cells (Table 1). Spheroidal neurons observed especially in transverse sections may correspond to the fusiform cells of sagittal and dorsal planes.
|Laminae||Transverse section||Sagittal section||Dorsal section|
|I||N = 502, 124.62 ± 49.75||N = 397, 112.43 ± 18.62||N = 95, 58.2 ± 17.3||N = 994, 113.10 ± 40.30||N = 104, 168.56 ± 45.9||N = 497, 135.28 ± 55.14||0||N = 601, 140.83 ± 51.16||N = 1,253, 101.55 ± 20.07||N = 731, 83.24 ± 35.93||0||N = 1,984, 94.8 ± 27.55|
|II||N = 1,798, 99.21 ± 52.66||N = 1,312, 67.75 ± 18.64||N = 903 59,3 ± 12,58||N = 4,013, 79.9 ± 40.3||N = 203, 177.3 ± 26.5||N = 731, 104.7 ± 11.70||0||N = 934, 117.9 ± 37.27||N = 1,198, 106.96 ± 16.3||N = 823, 88.63 ± 8.93||0||N = 2,021 98.81 ± 16.7|
|III||N = 311, 136.04 ± 41.82||0||0||N = 311, 136.4 ± 41.2||N = 183, 140.6 ± 76.5||0||0||N = 183, 140.96 ± 76.25||N = 214, 233.33 ± 25.59||0||0||N = 214, 233.33 ± 25.59|
|IV||N = 298, 99.58 ± 42.13||0||0||N = 298, 99.8 ± 42.3||N = 174, 108.6 ± 11.4||0||0||N = 174, 108.06 ± 11.24||N = 221, 151.63 ± 43.52||0||0||N = 221, 151.63 ± 43.52|
|V–VI||N = 306, 210.67 ± 40.51||0||0||N = 306, 210.67 ± 40.51||N = 488, 174.47 ± 74.4||N = 201, 259.59 ± 146.23||0||N = 689, 186.63 ± 105.31||N = 478, 219.45 ± 169.62||N = 98, 187.94 ± 105.71||0||N = 576, 214.2 ± 152.26|
|VII||N = 1,645, 340.29 ± 91.6||0||0||N = 1,645, 340.29 ± 91.6||N = 2,137, 338.69 ± 112.||0||0||N = 2,137, 338.69 ± 112.4||N = 2,322 375.15 ± 130.02||0||0||N = 2,322, 375.15 ± 130.02|
|VIII||N = 1,125, 597.87 ± 152.12||0||0||N = 1,125, 597.87 ± 152.12||N = 805, 406.09 ± 146.75||N = 723, 352.54 ± 175.83||0||N = 1,528, 381.1 ± 157.42||N = 415, 269.55 ± 81.68||0||N = 421, 234.23 ± 63.05||N = 836, 248.36 ± 69.03|
|IX||N = 1,237, 780.34 ± 244.09||N = 195, 550.29 ± 107.74||0||N = 1,432, 751.59 ± 242.04||N = 349, 679.05 ± 472.62||N = 512, 595.94 ± 250.88||0||N = 861, 633.59 ± 343.78||N = 527, 213.65 ± 97.92||N = 195, 125.21 ± 30||0||N = 722, 188.38 ± 91.68|
|X||N = 513, 275.22 ± 42.63||0||0||N = 513, 275.22 ± 42.63||N = 247, 306.36 ± 71.25||N = 128, 169.33 ± 24.61||0||N = 375, 237.85 ± 90.29||N = 835, 354.06 ± 73.6||N = 872, 312.97 ± 100.78||0||N = 1,707, 333.51 ± 87.85|
A few neurons could be found in Laminae III and IV. In transverse, sagittal, and dorsal sections these cells exhibited a multipolar morphology with 3–4 primary dendrites (Fig. 4A,B; Table 1). Cells located in Lamina III were usually larger than those of Lamina IV (Table 1).
In the dorsal horn, the neuropil contained numerous dendritic profiles, puncta, and axonal-like processes, especially in superficial layers of the dorsal horn. Similarly, the highest diffuse neuropil staining was observed in Layers I and II (Fig. 3A–D).
Numerous darkly-stained nitrergic neurons with a multipolar morphology were found in the intermediolateral cell column of the thoracic and first lumbar segments of the spinal cord. These cells had a large (Table 1) and angular somata with 3–6 primary dendrites (Fig. 5A–D). In sagittal and dorsal sections, these cells were organized in discrete groups and emanated numerous processes that interconnected the same groups along the longitudinal axis of the spinal cord (Fig. 5E–H). From L14 to Ca1 there was a small group of neurons that lay within the lateral border of Lamina VII. This nucleus contained small neurons with triangular somata emanating 3–6 primary dendrites. Throughout the entire craniocaudal extension of the spinal cord, few multipolar neurons were seen in middle and medial portions of Lamina VII. Labeled dendrites, axons, and puncta were observed in the neuropil (Fig. 5A–H).
Ventral Horn (Laminae VIII, IX)
The ventral horn exhibited a moderate density of labeled cells in Laminae VIII and IX. The neurons located in Lamina VIII appeared multipolar, fusiform (especially in sagittal sections), and spheroidal (only in dorsal sections) (Fig. 6A–E; Table 1). The primary dendrites of fusiform neurons, originating from the opposite poles of the ovoidal somata, were especially directed dorsally and ventrally (Fig. 6A–D). The mean cross-sectional somal area of multipolar neurons was higher than that of fusiform cells (Table 1).
Multipolar and fusiform neurons could be observed also in Lamina IX. As in Layer VIII, these neurons exhibited a large somata (Table 1). The somatic motoneurons located in Lamina IX remained unstained (Fig. 6F). Neuropil showed multiple puncta distributed around the large somata of presumptive unstained somatic motor neurons (Fig. 6F).
Central Cord (Lamina X)
A prominent group of nitrergic neurons was uniformly distributed around the central vein that substitutes the central canal of the spinal cord (Fig. 7A). These cells showed a multipolar or bipolar morphology. Multipolar neurons, observed in every plane of section, had a polygonal somata that gave rise to 3–5 thin primary dendrites (Fig. 7B,C). Sagittal and dorsal sections revealed rostro-caudal oriented bipolar neurons with a fusiform cell body and two primary dendrites (Fig. 7D). The mean perikarial areas of multipolar and bipolar neurons are reported in Table 1. Lamina X contained a dense array of labeled dendrites; in addition, axon-like processes could be observed (Fig. 7A–D).
Few nitrergic axons were observed in dorsal and ventral roots. Axons coursing in Lissauers's tract and the dorsal funiculus appeared stained at all spinal levels. Interestingly, nitrergic fibers projecting from the intermediolateral cell column laterally into the white matter could be observed.
In the present study, the distribution and morphology of nitrergic neurons in the bottlenose dolphin spinal cord has been revealed using histochemical (NADPH-d) and immunohistochemical (nNOS immunoreactivity) procedures. As reported in previous studies, the NADPH-d reaction and the mouse anti-nNOS marked the same neurons (Dawson et al., 1991; Vincent and Kimura, 1992; Giraldi-Guimarães et al., 1999; Bombardi et al., 2011, 2006). In the bottlenose dolphin, the laminar distribution of the spinal nitrergic neurons is similar to that reported in mouse, rat, rabbit, dog, cat, and Primates, with somata localized with a high density in Laminae I, II, VII (intermediolateral cell column and nucleus intermediolateralis sacralis of terrestrial Mammals), and X (Brüning, 1992; Dun et al., 1992, 1993; Valtschanoff et al., 1992a, 1992b; Spike et al., 1993; Terenghi et al., 1993; Saito et al, 1994; Vizzard et al., 1994; Wetts and Vaughn, 1994; Pullen and Humphreys, 1995; Marsăla et al., 1998, 1999; Kluchová et al., 2000). Nevertheless, there are some differences in the distribution of spinal nitrergic neurons between the bottlenose dolphin and terrestrial mammals: (a) in the bottlenose dolphin the nitrergic neurons appeared more homogenously distributed throughout the different spinal levels; (b), there are more neurons in Laminae III and IV in terrestrial mammals than in the bottlenose dolphin; (c), the somatic motor neurons of the bottlenose dolphin were completely unstained. References to these latter elements are equivocal, because in terrestrial mammals, even within the same species, the somatic motor neurons appeared either to contain (Terenghi et al., 1993; Pullen and Humphreys, 1995; Marsala et al., 1998) or not (Bruning, 1992; Dun et al., 1992, 1993; Valtschanoff et al., 1992a; Spike et al., 1993; Vizzard et al., 1994; Saito et al., 1994; Wetts and Vaughn, 1994; Marsala et al., 1999) NADPH-d or nNOS. NADPH-d and nNOS stained axons were seen running across the Lissauers's tract and the dorsal funiculus. Since many nitrergic neurons has been found in the sensory ganglia (Bombardi et al., 2011), these axons may be primary afferent fibers.
NO acts as a diffusible gaseous molecule able to induce biological activity on adjacent cells located up to 100 µm from its site of release. The principal effect of NO is to stimulate cytoplasmic guanylyl cyclase (sGC) to synthesize cGMP (Wood et al., 1990; Garthwaite, 1991). Thus this molecule can modulate a broad range of neurophysiological processes, acting on different neuronal populations. The present work identifies a specific histochemical (NADPH-d) and immunohistochemical (nNOS-immunoreactivity) staining in the gray matter of the spinal cord of the bottlenose dolphin. In particular, the distribution and the morphological features of nitrergic spinal neurons suggest that NO may be involved in sensory processing, autonomic neurotransmission, and motor pathways regulation.
The nitrergic neurons located in the dorsal horn were localized mostly in Laminae I and II, where they were embedded in a dense neuropil staining. These results suggest that the nitrergic network of the dorsal horn is prevalently involved in pain sensory processing. Accordingly, the bipolar nitrergic cells with dendrites elongated along the rostro-caudal axis of the spinal cord resembled islet cells, which act as interneurons modulating the nociceptive information directed to the spinal cord. In agreement with the previous data, pharmacological studies revealed that different spinal functional processes, such as nociceptive transmission, central sensibilization, and hyperanalgesia, require the production of NO (Meller et al., 1992; Meller and Gebhart, 1993; McMahon et al., 1993). In the bottlenose dolphin, the involving of NO on nociceptive transmission is also suggested from a recent study indicating that the majority of nitrergic neurons of the spinal ganglia express Substance P (SP) (Bombardi et al., 2011).
The distribution of nitrergic cells in the spinal cord has been often related to the autonomic preganglionic neurons (Dun et al., 1992, 1993; Valtschanoff et al., 1992a; Saito et al., 1994; Vizzard et al., 1994; López-Figueroa et al., 1996, 1997; Marsăla et al., 1998, 1999). In the present study, nitrergic and presumably autonomic neurons were observed in the intermediolateral cell column. These data, and the presence of stained axons only in the thoracic and upper lumbar ventral roots, suggest that NO may be involved especially in transmission of sympathetic ganglia. The small nitrergic nucleus located at the lumbar-caudal (L14-Ca1) level might be homologous to the parasympathetic nucleus intermediolateralis sacralis of terrestrial mammals. Our results suggest that in Cetacean, as in other mammals, NO contributes to the regulation of autonomic functions, in addition to the classical transmitters acetylcholine and norepinephrine. Accordingly, nNOS activity has been demonstrated in central and peripheral sites involved in autonomic regulation (Bredt et al., 1990; Patel et al., 2001; Toda and Okamura, 2003). Functional studies demonstrated that spinal NO enhances both excitatory and inhibitory neurotransmission in the intermediolateral cell column (Wu and Dun, 1995, 1996; Wu et al., 1997). Moreover, NO, located in sympathetic nerves, can be released as a co-transmitter in the peripheral autonomic system and induces vasodilator effects (Toda and Okamura, 2003). It is well known that Cetaceans have morphophysiological features that permit them to live in aquatic environment (Nagel et al., 1968; Vogl and Fisher, 1981; Ochrymowych and Lambertsen, 1984; Melnikov, 1997; Bombardi et al., 2010, 2011). In particular, during diving, blood is shifted to the huge retia mirabilia, to allow physiological distribution of fluids, permit thoracic compression at depth, heat the spinal cord, and regulate the oxygenation of the brain during the long breath-holding dives. Different studies have indicated that nNOS-derived NO is involved in local regulation of vascular tone acting through direct or indirect effects on vascular smooth muscle (Toda and Okamura, 2003; Hatanaka et al., 2006). Interestingly, the vascular smooth muscle of perispinal retia mirabilia is innervated by fibers containing nNOS-immunoreactivity (personal observations). Since previous studies have demonstrated that nNOS-derived NO could be produced by autonomic fibers and capsaicin-sensitive sensory nerve fibers (Toda and Okamura, 2003; Hatanaka et al., 2006), we could retain that NO acting on vascular tone of retia mirabilia may be produced by peripheral autonomic nerve fibers and primary afferent fibers. Accordingly, Bombardi et al. (2011) showed that in the spinal ganglia the majority of nNOS-IR neurons expressed SP. On the basis of the previous data, NO may be crucial to regulate the vascular tone of the retia mirabilia during diving. The increasing environmental pressure encountered during descent promotes compression of the relatively flexible thorax. The simultaneous shift of thoracic blood into the spinal retia mirabilia is one of the key factors that avoid fluid-induced damages to the heart and lungs. Experimental compression of the rat spinal cord demonstrated that NO dilates the spinal arterioles (Ishikawa et al., 2002). The same mechanism may induce nitrergic cells to adjust vascular caliber in the perispinal vessels of dolphins during diving.
The present study indicates that NO could be produced also by parasympathetic preganglionic spinal neurons. Potential visceral regulatory roles for the NO in these neurons could be similar to those of the intermediolateral cell column. A modulatory role of NO on parasympathetic activity was suggested by Toda and Okamura (2003) who identified nitrergic cells in parasympathetic ganglia that innervate the genitourinary system. In the bottlenose dolphin, however, the true nature of the role of NO systems on sympathetic and parasympathetic outflows remains to be fully explored and examined.
Lamina X receives numerous visceral afferents and it is probably involved in the transmission of visceral nociception. Numerous nitrergic neurons were observed in Lamina X. This aspect suggests a possible role for NO in visceral sensory processing, including visceral nociception.
Nitrergic neurons distributed in Laminae VIII and IX had the morphological features of interneurons. Thus, NO may regulate the transmission of motor signals using presynaptic mechanism.