Immunohistochemical investigations of the autonomous innervation of the cervine testis

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Immunohistochemical investigations of the autonomous innervation of the cervine testis Karl-Heinz Wrobel and Eveline Schenk Institute of Anatomy, University of Regensburg, Universit/itsstraBe 31, D-93040 Regensburg, Germany

Summary. The innervation of the cervine testis was studied in 6 roe deers, 7 red deers and 14 fallow deers. The results for the three species are rather similar. With antisera to neurofilament (NF) and neuron specific enolase (NSE), all small and large nerve fascicles can be demonstrated, but single fibers are incompletely stained. Immunoreactions against protein gene product-9.5 (PGP-9.5) and GAP-43 (growth-associated protein-43) are better suited to depict the complete innervation pattern. Bundles of the superior spermatic and inferior spermatic nerves reach the testis via three access routes as funicular, mesorchial and caudal nerve contributions. We found no morphological evidence that the nerves in the cervine testis are directly involved in regulating Leydig cell function or seminiferous tubular motility. The majority of the testicular nerves are associated with the testicular arteries, but the musculature in the walls of the venous plexus pampiniformis is also innervated. All vascular nerve fibers represent postjunctional sympathetic axons displaying a strong dopamine-beta-hydroxylase (DBH) activity, mostly co-expressed with neuropeptide Y (NPY). The presence of cholinergic fibers in the testis of the deer is only sporadic and probably of no functional importance. In all three species of deer, a small quantity of myelinated nerve fibers is encountered in spermatic cord and tunica albuginea and regarded as afferent. The viscerosensory quality in the testicular intrinsic innervation is very likely mediated by the CGRP (calcitonin gene-related peptide)positive fibers that run independently from the testicular vessels and end in the connective tissue of spermatic cord and tunlca albuginea. The testis of the red deer contains significantly more VIP (vasoactive intestinal polypeptide)-positive axons than that of roe and fallow deer. The

nerve density in the interior of the testicular lobules shows no regional differences, but there are age- and season-related changes that correlate with the developmental and functional state of the seminiferous tubules. Small testes with solid and narrow tubules, as seen in the prepuberal phase and during seasonal reproductive quiescence, are better innervated than large testes with expanded and spermatogenetically active seminlferous tubules.

Key words: Testis - Innervation - Red deer - Fallow deer Roe deer - Immunohistochemistry

Introduction

The innervation of the mammalian testis and epididymis is effected by the superior and inferior spermatic nerves (SSN and ISN) which both contain efferent and afferent fibers. The efferent fibers in the SSN arise from the intermesenteric and renal plexuses and descend along the testicular artery to the cranial pole of the testis; the bundles of the ISN also contain fibers from the pelvic plexus and accompany the ductus deferens (Kuntz and Morris 1946; Hodson 1970). The dorsal root ganglia innervating the testis via the SSN are localized in the first lumbar segments and more important for the afferent visceral nervous input from the scrotal contents than the afferent fibers in the ISN that have their perikarya in dorsal root ganglia of the sacral region (Tamura et al. 1996). A certain percentage of the efferent fibers to the testis come from plexuses of both the ipsilateral and the contralateral sides (Rauchenwald et al. 1995). Also a small fraction (about 8%) of the afferent fibers within the SSN travel to Correspondence to: K.-H Wrobel dorsal roots of the opposite side (Kuo et al. 1983; Tamura Fax: +49 9419 43 28 40 E-mail: [email protected] et al. 1996). The findings of a contralateral or bilateral inAnn Anat (2003) 185:493-506 © Urban & FischerVerlag

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nervation of the testis may help to explain pathophysiological bilateral consequences following unilateral testicular torsion, cryptorchidism or experimentally induced varicocele (Horica et al. 1982; Green et al. 1985; Chakraborty et al. 1985; Anderson and Williamson 1986; Rauchenwald et al. 1995; Tarhan et al. 1996; Collin et al. 1998). Though the functions of the testis are primarily under hormonal control, the autonomous nervous system also plays an important role in regulating the activities of the male gonad and its excurrent duct system. In all species studied so far, the intrinsic testicular innervation is involved in the control of blood flow and intrascrotal temperature. In addition, however, there may be a direct or indirect influence of the nervous system on gonadotropin secretion, testosterone production and spermatogenesis. Experimental data furnish evidence that there exists a nervous circuit between brain and testis that directly regulates gonadotropin release and testosterone production (Gerendai and Halasz 1981; Mizunuma et al. 1983; Preslock and McCann 1985; Chow et al. 2000; Lee et al. 2002). In a number of mammalian species, intimate topographical relationships between Leydig cells and testicular nerves have been observed (Yamashita 1939; Okkels and Sand 1940; Gray 1947; Van Campenhout 1949; Stach 1963; Baumgarten and Holstein 1967; Norberg et al. 1967; Nistal et al. 1982; Prince 1992; Wrobel and Brandl 1998; Feldmeier 1998; Wrobel and Gtirtler 2001). Noradrenaline receptors have been localized on the surfaces of Leydig cells in mouse, rat and hamster (Anakwe and Moger 1984; Anakwe et al. 1985; Mayerhofer et al. 1989) and stimulation of the SSN increases testosterone secretion in cats (Chiocchio et al. 1999). A participation of the nervous system in the intratesticular transport of spermatozoa is still a matter of debate. In the human testis, non-myelinated fibers have been described close to the myofibroblast layer within the tubular lamina propria (Nistal et al. 1982; Yamamoto et al. 1987), though Prince (1996) could demonstrate such tubular fibers only in the immature testis during childhood. From pharmacological experiments with rat seminiferous tubules, Miyake et al. (1986) conclude that their contractions elicited by excitation of adrenergic nerves may be an important factor for intratesticular sperm transport. Pholpramool and Triphrom (1984), on the other hand, were unable to produce observable effects on rat seminiferous tubules by application of a number of cholinergic and adrenergic agonists. A participation of the nervous system in the extrusion of spermatozoa from the seminiferous tubules is more likely in species with smooth muscle cells in their testicular capsule, viz. rat, dog, rabbit (Davis and Langford 1971; Bell and McLean 1973; Leeson and Cookson 1974) or donkey (Wrobel and Moustafa 2000). In a series of recent studies, we have investigated the innervation of the testis in a number of mammalian species with a similar methodology: bull (Rose 1992; Wrobel and Abu-Ghali 1997; Abu-Ghali 1997), pig (Wrobel and Brandl 1998; Brandl 1998), donkey (Wrobel and Moustafa 2000), cat (Wrobel and Gtirtler 2001; Gtirtler 2001),

camel (Saleh et al. 2002 a, b) and man (Feldmeier 1998) and noted considerable species-specific variations in report to nerve density, distribution pattern and transmitter content. These variations are particularly obvious in the group of ungulates when members of the different families (Equidae, Suidae, Bovidae, Camelidae) are compared. In the present investigation, we extend our results by describing the intrinsic testicular innervation in three species of deer, representing yet another family of artiodactyls (Cervidae).

Material and methods This investigation was carried out on the testicles of 6 roe deers, 7 red deers and 14 fallow deers. The material was collected in different seasons of the year and comprised gonads with seminiferous tubules in various states of spermatogenetic activity, When the testes were removed from their envelopes, the adjoining portion of the spermatic cord was preserved. Tissue samples of suitable size were taken from spermatic cord and different regions of the testis in a manner similar to previous studies on other species (see Wrobel and Moustafa 2000). Generally, one to two samples from each spermatic cord and up to 18 samples per testis were collected in order to recognize the complete innervation pattern. Immersion fixation was carried out in two steps. Fixative I (30rain) contained 4% paraformaldehyde; 15% v/v saturated picric acid; 0.1% glutaraldehyde in 0.1M phosphate buffer, pH 7.4. Fixative II (several hours) had the same composition as fixative I but without glutaraldehyde. Following fixation, the blocks were washed in 0.1 M phosphate buffer, transferred into a graded series (10%, 20%, 30%) of saccharose-eontaining rinsing buffer. Then the samples were immersed in Tissue Tek OCT compound (Miles, Elkhardt, Ind., USA) and snap-frozen in liquid nitrogen. Cryostat sections (12 pm thick) were mounted on gelatin/chrome alum-coated slides and air-dried for 2-3 rain before further treatment.

Acetylcholinesterase(ACHE)histochemistry For AChE histochemistry the modified, direct-colouring method of Kujat et al. (1993) was applied. Following a brief incubation period (1-2 h), this method allows for the specific visualization of cholinergig nerves comparable to the results of cholinacetyltransferase (ChAT)-immunoreactivity. The modified AChE method has the following steps: (1) Rinsing and preincubation at 4 °C for 10 min in solution I: 0.05 M morpholinoethane (MES), pH 5.5 (Sigma, Mtinchen, Germany); 0.01 M citrate buffer, pH 5.5; Karion F diluted 1:5 (Merck, Darmstadt, Germany); 10-4 M Iso-OMPA (K & K/ICN, New York, USA). (2) Incubation (4 °C) for 1-2 h or 24 h, respectively in the substrate medium: 0.05 M MES, pH 5.5; 0.01 M citrate buffer, pH 5.5; Karion F diluted 1 : 5; 0.003 M CuCI2; 0.005 M acetylcholine chloride (Sigma); 0.0005 M K3Fe(CN)6; 10-4 M IsoOMPA. (3) Rinsing in solution II (as solution I, but without Iso-OMPA) for 10 min. (4) Dehydration and embedding in DPX. Controls included: (a) Omission of the substrate from the incubation medium. (b) Inhibition of the reaction by 10-2 M diisopro-

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pylfluorophosphate (Serva, Heidelberg, Germany) instead of Iso-OMPA in rinsing solution I and incubation medium. (c) Inhibition of the reaction with 5 x 10-5 M BW 284c51 (Sigma) added to rinsing solution I and incubation medium. The results of all these controls were negative. As a positive control tissue, bovine M. retractor penis was used.

Results C o m p a r i s o n b e t w e e n various pan-neuronal markers

For the demonstration of the autonomous innervation of cervine testis and epididymis four different pan-neuronal Immunohistochemistry markers were used and compared, namely immunoreacAll subsequent steps were carried out in a moist chamber on tions to NF, NSE, P G P 9.5 and GAP-43. Techniques using slides with sections surrounded by water-repellent PAP-PEN antisera to NF and NSE completely depict all larger (SCI Science Services, Mtinchen, Germany): nerve bundles and fascicles, but fail to stain isolated single axons and the entirety of the intramural vascular (1) Preincubation (60min) with blocking buffer containing plexuses. Demonstration of the nervous components with 0.1 M TRIS (pH 7.4); 0.15% Thimerosal; 0.8% Triton X100; 0.8% NaC1; 20% normal goat serum (DAKO, Ham- antiserum against P G P 9.5 allows the innervation pattern burg, Germany); 20% foetal calf serum (Seromed, Mttn- to be followed from the large bundles to the finest terminal ramifications and was therefore selected as the methchen, Germany). (2) Rinsing (3x10 min) in TBS: 0.1 M TRIS (pH 7.4); 0.8% od of choice for the description of the general nerve NaC1; 0.0015% Triton X-100. distribution, in spite of an unwanted staining in several (3) Incubation (overnight) with primary antibody in blocking other testicular components. The immunoreaction to buffer at room temperature: GAP-43 is also well suited to demonstrate the overall in• 1:1000 mouse-anti-neurofilament (NF-200); s o u r c e : Signervation pattern in the deer testis. All fibers innervating ma-Aldrich, Deisenhofen, Germany. the arteries in the cervine tissue samples are DBH-posi• 1 : 5000 rabbit-anti-human protein-gene-product-9.5 (PGPtive. Since in the D B H sections a background staining is 9.5); s o u r c e : UltraClone, Isle of Wight, UK. • 1:5000 rabbit-anti-human neuron specific enolase (NSE); virtually absent, this specific reaction can favorably supplement the results for the vascular innervation obtained source: Cambridge Research Biochemicals, Cambridge, with the pan-neuronal markers. UK. • 1 : 100 mouse-anti-rat growth-associated protein-43 (GAP43); source: Chemicon Internat. Inc., Temecula, CA, USA • 1:1000 rabbit-anti-human myelin basic protein (MBP); A c c e s s routes o f the intrinsic nerves to the cervine testis source: DAKO Diagnostica, Hamburg, Germany. • 1 : 1000 rabbit-anti-bovine dopaxninc-beta-hydroxylase(DBH); The nerve distribution in the testis of the three deer spesource: ETI, Allendale, NJ, USA. • 1:100 rat-anti-mouse tyrosine hydroxylase (TH); source: cies studied is very similar and therefore reported together in a general manner. Separate results for fallow, Boehfinger, Germany. • 1:5000 rabbit-anti-swine neuropeptide Y (NPY); source: red, and roe deer are mentioned only when characteristic Cambridge Research Biochemicals. species-specific differences are encountered. • 1 : 5000 rabbit-anti-swine vasoactive intestinal polypeptide The nerve fibers of the SSN and ISN reach their targets (VIP); s o u r c e : Cambridge Research Biochemicals. via three different access routes. (1) The funicular or cra• 1:2000 rabbit-anti-rat calcitonin gene-related peptide nial nerve contribution is part of the SSN and represented (CGRP); source: Cambridge Research Biochemicals. by nerve bundles inside the vascular cone of the sper• 1:2000 rabbit-anti-substance P (SP); source: Eugene Tech matic cord. The vascular cone is composed of the coiled International Inc., Ridgefield Park, NJ, USA. testicular artery with its supratesticular branches and of (4) Rinsing as in step 2. (5) Incubation (60 min) in secondary antibody/biotinylated in the venous plexus pampiniformis and covers the cranial pole of the testis. (2) The mesorchial nerve contribution blocking buffer. (6) Rinsing as in step 2. reaches the testis by traversing the mesorchium. (3) The (7) Blocking of endogeneous peroxidase with phenylhydrazine. caudal nerve contribution is part of the ISN, accompanies (8) Rinsing in TBS. the ductus deferens to the epididymal border and is (9) Incubation (60 rain) in AB complex (ABC); source: Vector, primarily involved in the innervation of the epididymal Burlingame, CA, USA. tail region. A smaller number of fibers, however, travel (10) Rinsing as in step 2 and developing with 0.5 mg/ml DAB; through the ligamentous bridge between testis and epidi(source: Sigma) in 0.1 M TRIS (pH 7.4) containing 0.002% dymal tail (Lig. testis proprium) to the caudal pole of the COC12 ( 6H20; 0.4% • 6H20 and 0.012% H202. gonad. (11) Rinsing once in TBS, dehydration, mounting in DPX. C o n t r o l s f o r i m m u n o h i s t o c h e m i s t r y . Controls included: (a) Omission of the primary antiserum. (b) Substitution of primary antiserum by non-immune serum 1 : 500 in blocking buffer. (c) Blocking of the primary antibody by preincubation with the matching antigen in excess. No immunostaining was observed after any of these control procedures (a-c).

N e r v e distribution in the vascular cone and testis

The vascular cone of the spermatic cord contains a number of thick nerve bundles (Fig. 1) which accompany the testicular artery to the cranial testicular pole. Whereas

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Fig. 1. Fallow deer, spermatic cord, NF-IR, x 140. A large branch of the SSN is heavily stained, whereas the nerve plexus in the wall of the A. testicularis (a) is incompletely depicted by this reaction. Hg. 2. Red deer, spermatic cord, PGP-9.5-IR, x225. The intramural nerve plexus of the A. testicularis (a) is situated at the adventitia-media border. Bar in fig. 2 also applies to figs. 3-6. Fig. 3. Red deer, spermatic cord, DBH-IR, x225. This longitudinal section through the wall of the A. testicularis reveals an unequal density of the intramural nerve plexus. Fig. 4. Red deer, spermatic cord, DBH-IR, x 225. Nerve supply of the venous pampiniform plexus. Fig. 5. Roe deer, tunica albuginea, DBH-IR, x 225. Relatively loose nerve plexus in two coils of a large tunical artery (a). Fig. 6. Red deer, tunica albuginea, GAP-43-IR, x 225. Smaller tunical arteries possess a dense intramm'al nerve plexus. Compare to Fig. 5.

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Fig. 7. Red deer, septulum testis, NF-IR, x 225. Thick nerve bundles accompany a septular artery (a) on its way to the centrally located mediastinum testis. Bar in fig. 7 applies also to figs. 8 and 11. Fig. 8. Red deer, septtflum testis, GAP-43-IR, x 225. Note the dense and regular innervation of a septular artery (a). Fig. 9. Red deer, mediastinum testis, NF-IR, x 90. Nerve bundles of the funicular contribution perforate the tunica albuginea at the cranial testicular pole and are then located at the outer periphery of the mediastinum testis (m). Fig. 10. Fallow deer, DBH-IR, x 140. The nerve bundles (arrowheads) at the border between mediastinum (m) and testicular parenchyma (p) supply the dense nerve plexuses around the arterial convolutes (arrows) and also dismiss fibers to the small arteries in the interior of the mediastinum. Fig. 11. Fallow deer, DBH-IR, x 225. Nerves surround the terminal segments (ts) of the seminiferous tubules in the spermatogenetically active fallow deer.

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the testicular artery is increasingly coiled, the nerve bundles take a straight or slightly undulating course within the venous network of the pampiniform plexus. From these large nerve bundles, smaller branches or single fibers deviate into the perivascular stroma and innervate the walls of testicular artery, epididymal arteries, small nutritive arteries, plexus pampiniformis and the stroma itself. Within the arterial walls, the vascular nerves form a network at the media-adventitia border. In the A. testicularis, this plexus is relatively loose and of variable density (Figs. 2, 3); in the smaller arteries, the intramural arterial nerve plexus is much denser and equally present in the whole circumference of the vessels. The innervation of the venous walls in the pampiniform plexus (Fig. 4) is generally weaker than the arterial innervation. Single fibers form a loose network in the periphery of the veins, but some of them also invade the media and end between the muscle cells. Furthermore, the nerve distribution in the pampiniform plexus displays regional differences in density which are particularly obvious in the red deer, where the frequency of nerves increases towards the cranial testicular pole. The nerve bundles to the testis enter the tunica albuginea via the funicular, mesorchial and caudal access routes. Nerves of the funicular contribution concentrate at the epididymal border, but are also present at the ventral and lateral sides of the testis. They run slightly obliquely to the longitudinal axis of the testis in caudal direction. The mesorchial set of tunical nerves is the smallest and consists of a number of single fibers and thin bundles that enter the tunica at the epididymal side and follow more a circumferential course. The nerves of the caudal access route pass through the narrow ligamentous bridge to the epididymis and distribute within the tunica of the caudal testicular pole where they diverge in cranial direction. By this arrangement of the tunical nerves, their course is different from that of the intratunical arteries. Where both structures come in close proximity to or cross each other, small nerve bundles deviate towards the arteries and supplement the intramural arterial plexus which is dense in such sites of mutual contact and looser in the distances in between (Figs. 5, 6). The thin-walled veins of the stratum vasculare in the tunica albuginea and their feeder veins in septula and lobuli testis are generally not innervated at all. When the intratunical arteries and nerve bundles enter the so-called septula testis, which in the deer testis are strands of connective tissue, both structures now run parallel and in close proximity to each other towards the centrally located mediastinum testis (Fig. 7). Septular arteries possess a dense and regular nerve plexus in their walls (Fig. 8). They end in the periphery of the mediastinum in form of characteristic arterial convolutes which are also heavily innervated (Fig. 10). In the caudal half of the testis, the nerves to the mediastinum exclusively come from the terminal ramifications of the septular nerve bundles. In the cranial half of the testis, most mediastinal nerve bundles are branches

of the funicular contribution which perforate the tunica albuginea at the cranial testicular pole. On their way to the equatorial region of the testis, these bundles travel into the periphery of the mediastinum (Figs. 9, 10) and continously dismiss fibers into the interior of the mediastinum, but also to the arterial convolutes in their neighborhood. In the interior of the mediastinum, single nerve fibers accompany and innervate the small mediastinal arteries, and are also observed free in the mediastinal stroma, but make no contacts with the epithelium of the rete testis. The testicular parenchyma proper of the cervine testis (seminiferous tubules and Leydig cells) is not innervated. The intralobular arteries and arterioles, however, are supplied by vascular nerves. Some individual axons are observed in the connective tissue between the tubules, but their occasional topographic proximity to Leydig cells and tubular walls seems to be merely accidental. In samples of adult fallow deer, a loose nerve plexus surrounds the terminal segments of the seminiferous tubules and their transition into the straight testicular tubules (Fig. 11). The vascular nerves in the interior of the cervine testis show differences depending on the functional state of the germinal epithelium (Figs. 12, 13). In animals with solid tubules and small tubular diameters, a situation typical for prepuberal specimens and older animals during the phase of annual quiescence, the testicular nerve density is high. All intralobular arteries possess a dense, continuous intramural nerve plexus as far as a multilayered musculature is present in their tunica media. The smaller arterioles with only one layer of musculature are accompanied by one or two individual nerve fibers. In animals with somewhat larger tubular diameters and beginning or decreasing spermatogenesis, the intramural nerve plexuses in the walls of the larger intralobular arteries are less dense and the smaller arterial branches between the tubules may not be innervated at all. In samples which display heavily coiled seminiferous tubules with maximal diameters, a spermatogenetically fully active tubular epithelium and expanded Leydig cell groups, the nerve density is low. Some of the septular arteries, most of the enlarged intralobular arteries and their smaller ramifications between the tubular coils are free from any innervation.

The differentiation of testicular nerves by specific reactions MBP-IR: A small percentage of fibers in the nerve bundles observed in spermatic cord and tunica albuginea in most samples of all three species of deer reveal a positive reaction (Figs. 14, 15). These myelinated fibers are never seen in the interior of the gonad, but end close to the surface of the testis. D B H - I R , TH-IR: Both reactions are characteristic for postjunctional sympathetic fibers, but T H - I R stains signif-

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16 Fig. 12. Fallow deer, DBH-IR, × 225. In a small testis with solid seminiferous tubules a high nerve density is encountered. Fig. 13. Roe deer, DBH-IR, × 225. In a large testis with spermatogenetically active seminiferous tubules a low nerve density is present. Compare to fig. 12. Bar in fig. 13 applies also to figs. 12, 16, 17. Fig. 14. Fallow deer, tunica albuginea, MBP-IR, x 350. A nerve bundle contains a number of myelinated fibers in regular distribution. The black structure indicated by arrow is a small blood vessel with positive erythrocytes. Bar in fig. 14 applies also to fig. 15. Fig. 15. Fallow deer, tunica albuginea, MBP-IR, × 350. A solitary myelinated nerve fiber displays a node of Ranvier (arrow). Fig. 16. Red deer, septulum testis, AChE (2 h), × 225. One of the rare cholinergic fibers accompanies a septular artery (a). Fig. 17. Fallow deer, spermatic cord, NPY-IR, ×225. Most fibers in the thick bundles of the SSN and the majority of axons in the walls of the venous pampiniform plexus are positive.

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Fig. 18. Fallow deer, small testis with solid tubules, NPY-IR, × 225. Notice the dense plexus in the arterial walls and many intertubular fibers. Bar in fig. 18 also applies to figs. 19, 20, 22, 23. Fig. 19. Fallow deer, large testis with spermatogenetically active tubules, NPY-IR, × 225. Notice the loose plexus in the arterial wall (lower right) and sporadic fibers between the tubular coils (arrow). Fig. 20. Fallow deer, tunica albuginea, VIP-IR, ×225. Single positive fibers can be seen in the connective tissue and as constituents of a larger nerve bundle (arrow). Fig. 21. Red deer, VIP-IR, × 350. A positive varicose axon between seminiferous tubules. Fig. 22. Red deer, septulum testis, VIP-IR, x 225. The intramural plexus of a septular artery (a) contains many VIP-positive fibers. Fig. 23. Red deer, spermatic cord, CGRP-IR, × 225. Some positive fibers occur in the bundles of the SSN.

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icantly fewer axons than DBH-IR. Nearly all nerve fibers, reacting positively with the pan-neuronal markers, particularly all vascular nerves in the samples, are DBH-positive. Due to the intense staining of their varicosities, the terminal ramifications of the reacting nerve fibers often resemble a string of beads. AChE reaction: Our modified AChE reaction with short incubation (2 h) is specific for cholinergic fibers. A small contingent of them is found in the bundles of SSN and ISN, their main target being the wall of the ductus deferens. Sporadic positive axons are seen in the periphery of the mediastinum testis, in the septula testis (Fig. 16) and in the tunica albuginea of the caudal testicular pole. NPY-IR: Many fibers in the larger nerve bundles of the samples are positive for NPY, the other axons are frequently grouped together in negative sub-bundles (Fig. 17). Within the intramural arterial plexuses, nearly all axons present contain this neuropeptide. Particularly high NPY concentrations are encountered in the walls of the arterial convolutes, flanking the mediastinum. The nerve plexus surrounding the terminal segment of the seminiferous tubule in the fallow deer is made up of DBH- and NPY-positive fibers. Some NPY-positive nerves reach the walls of the venous pampiniform plexus (Fig. 17). Inside the testicular lobules, the nerve distribution varies according to the degree of tubular development. The innervation is denser and the NPY reaction is stronger in testes with solid tubules and small tubular diameters (Fig. 18); the innervation has thinned out and the single axons often react less strongly in testes with tubules that possess a distinct lumen and display ongoing spermatogenesis (Fig. 19).

VIP-IR: The nerve bundles of the ISN traveling with the deferent duct contain a certain number of VIP-positire axons, whereas the bundles of the SSN in the vascular cone and the nerves in the walls of A. testicularis and pampiniform plexus are VIP-negative. Most of the VIPIR axons in the ISN end in the mucosa and muscular coat of the deferent duct where they even occur in intraepithelial position. Others terminate around the loops of the epididymal tail, but some reach also the testis via the mesorchial and caudal access routes. In fallow and roe deer, the tunica albuginea contains small bundles and individual varicose axons which tend to take a circumferential course (Fig. 20). The nerve plexuses in the walls of the tunieal and septular arteries are generally negative. A small number of positive fibers are observed in the periphery of the mediastinum testis and also within the testicular parenchyma. The intraparenchymal fibers are situated between the seminiferous tubules (Fig. 21) and predominate in the subtunical area and the caudal half of the testis. The testis of the red deer contains significantly more VIP-positive nerves than the male gonad of fallow and roe deer. Many tunical and septular arteries have VIP-positive axons as components of their intramural plexuses (Fig. 22). In all testicular regions, the septular arteries can be accompanied by fine VIP-positive bundles, the fibers of which deviate into the lobules of the parenchyma. Intralobular fibers can be observed close to smaller arteries, but also apparently independent from blood vessels near tubular walls and within Leydig cell groups. CGRP-IR: Solitary CGRP-positive axons are observed as components of the bundles of both SSN (Fig. 23) and ISN, but more of the CGRP-positive fibers travel into the branches of the ISN. Small bundles in the neighborhood

Fig. 24. Red deer, spermatic cord, CGRP-IR, x 225. CGRP-positive fibers (arrows) are observed in the connective tissue of the vascular cone. Bar in fig. 24 also applies to fig. 25. Fig. 25. Red deer, tunica albuginea, CGRP-IR, x 225. Solitary positive fibers run independently from blood vessels in the superficial layer of the tunica albuginea. Arrow indicates testicular surface. 501

of the vas deferens and within the mesorchium may be positive in their entirety, indicating that the CGRP-IR axons are grouped together and separated from the rest. Most of the positive axons end within the stroma surrounding the vas deferens, in the wall of the A. ductus deferentis and between the coils of the epididymal duct, but some reach the testis proper via mesorchial and caudal access routes. Of the SSN, single fibers occur in the connective tissue between the veins of the pampiniform plexus (Fig. 24). In the fallow deer, the intramural nerve plexuses of A. testicularis and its smaller branches may display also some positive fibers. From the structures of the testis itself, the tunica albuginea contains a small quantity of CGRP-positive nerves which generally run independently of the arteries of the tunica vasculosa (Fig. 25). The interior of the testis (septula, lobules) is devoid of CGRP-IR axons. This is the case in the mediastinum testis in red and roe deer, whereas in the fallow deer, some solitary fibers may be present in the periphery of the mediastinum, but only in the cranial half of the gonad. SP-IR: The nerve bundles of the SSN are negative, those of the ISN occasionally display a few positive fibers which end in the walls of ductus deferens and epididymal tail. A rare SP-positive axon may occur in the connective tissue of the tunica albuginea (fallow deer) or the pampiniform plexus (red deer).

Discussion Similarly to the testes of the bull (Wrobel and Abu-Ghali 1997), pig (Wrobel and Brandl 1998), donkey (Wrobel and Moustafa (2000) and camel (Saleh et al. 2002 a), the branches of the cervine SSN and ISN reach the testis by three access routes. (1) The cranial or funicular nervous contribution is composed of bundles from the SSN only, which lie inside the vascular cone of the spermatic cord and approach the cranial pole of the testis. (2) The mesorchial nervous contribution carries fibers to the epididymal side of the testis. (3) The caudal nervous contribution crosses the ligamentous bridge between epididymal tail and the caudal testicular pole. We found no morphological evidence that the nerves in the cervine testis are directly involved in regulating Leydig cell function or seminiferous tubular motility. With the exception of the few myelinated fibers and some VIPand CGRP-positive axons, the vast majority of the cervine testicular nerves are associated with blood vessels, represent DBH-positive postjunctional sympathetic fibers and must therefore be regarded as primarily vasomotor in function. As in other mammalian species, the main target for the testicular vascular innervation is the smooth musculature in the arterial walls where the fibers form plexuses at the media-adventitia border. Of the testicular veins, normally only the elements constituting the pampiniform plexus are supplied by nerve terminals. In the camel, however, the complete venous side of the testicular

circulation, including pampiniform plexus, is devoid of intrinsic innervation (Saleh 2002 a), and in bull and cat the veins in the stratum vasculosum of the tunica albuginea also exhibit sporadic contacts to nerve fibers (Wrobel and Abu-Ghali 1997; Wrobel and Gtirtler 2001). The testis of the deer exhibits seasonal changes (Chapman and Chapman 1970: Chaplin and White 1972; Asher et al. 1987, 1989) in regard to testicular volume, seminiferous tubular diameter, spermatogenetic activity and the degree of intertubular (lobular) vascularization. These variations are accompanied by changes in intralobular nerve density. In animals with small and solid tubules, typically found during annual reproductive quiescence, but also around puberty, all intralobular arteries possess a dense and continuous innervation as far as their capillary transition. In the active season of the year, testicular volume and seminiferous tubular diameter are maximal, regular spermatogenesis takes place in all parts of the testis and the Leydig cell population is enlarged. In adaptation to these progressive features seen in the testicular parenchyma, the vascularization has also reacted: Existing arteries in the septula and between the tubules have grown in length, acquired larger diameters and more musculature in their media. In addition, new intertnbular vessels have apparently been formed by angiogenesis at the end of the arterial vascular tree. All these characteristics of an increased testicular function however, are not followed by a similar development of the intrinsic testicular innervation. On the contrary, in this activated functional state, most of the intralobular and intertubular arteries and even some of the septular arteries in all regions of the cervine testis are now free of any innervation. Similar changes in testicular innervation have been reported for the male camel (Saleh et al. 2002 a), an incomplete seasonal breeder. In this species, the gonad grows during the winter season to reach its greatest volume in spring. Then the testicular volume decreases again gradually during summer and autumn. The nerve content within the septula testis and in the intertubular region is lowest during winter and spring and increases during summer and autumn. At first glance, the findings in the deer and camel testes, seem to support the notion of Molenaar et al. (1997) and Sienkiewicz et al. (2000) who suggest an inverse relationship between innervation and endocrine activity of the testis. However, the coincidence of a low volume density, small individual cell sizes and a low 3-beta-HStDH activity of camel Leydig cells and the presence of very few intertubular nerves during the spring season (Saleh et al 2000 a) does not support the hypothesis of Molenaar et al. (1997). Also, the wavelike deployment of the porcine Leydig cell population seeri during normal postnatal development (Dierichs et al. 1973; Wrobel et al. 1973) is not inversely paralleled by the testicular innervation. Testes of piglets of three to five weeks exhibit the most intense testicular innervation of all age groups and at the same time a period of prepubertal Leydig cell hypertrophy (Wrobel and Brandl 1998).

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The age- and season-related changes in testicular nerve density that exist in most mammals correlate fairly well with the functional or developmental state of the seminiferous tubules. Testes with solid and small tubules as seen in the prepuberal phase of all mammals and during the periods of reproductive quiescence in seasonal breeders, are generally better innervated than those with expanded and spermatogenetically active seminiferous tubules. This suggestion is corroborated by several observations. Nistal et al. (1982) found more nerve contacts on human seminiferous tubules in children and in adults suffering from hypogonadism than in normal men. Prince (1996) could demonstrate the presence of nerve terminals in the human tubular lamina propria only during childhood (three to 10 years). Mayerhofer et al. (1996) observed abundant THpositive nerve fibers in the testes of juvenile rhesus monkeys, but only a few of them between the tubules of the adults. Particularly obvious is the retreat of the intrinsic innervation from the ungulate testis during the period of its impressive pubertal and postpubertal expansion. Whereas the testis of the immature piglet (three to five weeks) is still richly innervated, the extremely large testis of the adult boar (two to three years) is devoid of any intrinsic innervation (Wrobel and Brandl 1998). In the testis of the adult bull, the testicular parenchyma in the caudal half of the organ is no longer innervated (Wrobel and Abu-Ghali 1997), whereas in five to 25 week-old calves nerve fibers are present in this location. In the testis of the young-adult donkey (Wrobel and Moustafa (2000), the nerves have retracted in the direction of the two poles and the epididymal side and are completely absent from the large area between margo liber and the centrally localized mediastinum. The inhomogeneous innervation pattern in the interior of the testis and the retraction of the innervation to the sites from where the nerve fascicles have penetrated the organ, may indicate that in the young-adult donkey, the retreat of the nerves from the testis is still under way, concurrent with continuing testicular growth and seminiferous tubular expansion that take place through the first five years of life in this species (Nipken and Wrobel 1997). In the fully developed, spermatogenetically active testes of the fallow deer, a loose nerve plexus surrounds the terminal segments of the seminiferous tubules. The nerve fibers involved are DBH-positive. The transitional region between seminiferous and straight testicular tubules in mammals generally works in a valve-like manner, thus controlling the orthograde passage of tubulus seminiferus content and preventing a reflux from the rete testis (Tuck and al. 1970; Kormano 1974). At the level of the terminal segment, a regulation of the tubular lumen may be supposed, and in some mammals, special sphincter-like devices have actually been described. In the hamster, Cavicchia and Burgos (1977) report a ring of contractile cells at the border between the seminiferous and straight testicular tubules, and in the bull, each terminal segment is surrounded by.a cuff-like vascular plexus (Wrobel et al. 1978). The concentration of nerve fibers at this site in the testis of the fallow deer may also serve regulatory purposes.

The nerves that reach the mammalian testis are generally of the non-myelinated variety. However, in the large nerve bundles of the SSN in the supratesticular portion of the spermatic cord in cat (Peterson and Brown 1973) and camel (Saleh et al. 2002 a), a small number of weakly myelinated nerve fibers have been identified with the electron microscope. In the cat, these fibers amount to less than 3.6%, are afferent with a conduction velocity between 3.0 and 52.5 m/sec (Peterson and Brown 1973) and seem to terminate within the spermatic cord, since the MBP reaction is negative in the feline testis proper (Wrobel and Gt~rtler 2001). Also in all three species of deer, a small percentage of the fibers in the SSN is myelihated and MBP-positive. Such fibers are not restricted to the spermatic cord, but can be observed also in the tunica albuginea, close to the surface of the testis. They must be considered as afferent, though encapsulated corpuscular nerve terminals that have been described in the adult human tunica albuginea (Yamashita 1939; Kreutz 1964) are lacking in the cervine testis. The paucity of myelinated fibers in the mammalian testis seems to corroborate the notion that the senses of touch and pain in the gonad mainly involve the testicular coverings and their somatic innervation by the genitofemoral, pudendal, ilioinguinal and cutaneous femoris posterior nerves (Hodson 1970). On the other hand, the surgical treatment of chronic testicular pain requires, besides transsection of the ramus genitalis of the genitofemoral nerve, also stripping of the perivascular tissue of the testicular artery by bipolar diathermy (Choa and Swami 1992). This points to an involvement of visceral fibers in the SSN in the conveyance of orchialgia. The viscerosensory quality in the testicular intrinsic innervation is very likely mediated by SP-IR and CGRP-IR fibers, for both neuropeptides are consistent markers for sensory neurons in various parts of the body (Maggi 1991; Nishi et al. 2000). In the canine SSN, even a coexistence of CGRP and SP has been encountered (Tamura et al. 1996). CGRP is released from the endings of capsaicin-sensitive axons (Yamada et al. 1977; Ngassapa et al. 1998) and has important viscerosensory modulatory effects, but also a broad variety of other biological functions. In the three species of deer, a few CGRP-positive fibers are observed in the SSN, but significantly more lie within the bundles of the ISN. The majority of the positive axons terminate in the wall of the ductus deferens, between the loops of the epididymal canal and in the connective tissue of the pampiniform plexus. The CGRP-IR fibers that reach the confines of the testis proper via all of the three access routes distribute in the tunica albuginea, independently of the arteries of the tunica vasculosa. This localization is consistent with an assumed sensory function. Additionally, in the fallow deer the intramural nerve plexuses of the A. testicularis and its branches display some positive axons. Vascular CGRP is generally involved in a predominantly endothelium-independent vasodilation of blood vessels (Burgio et al. 1997; Lundgaard et al. 1997; Sheykhzade and Nyborg 1998; Yoshimoto et al. 1998). SP-positive fibers are absent in the

503

cervine SSN, but occur in the ISN; their destination is the wall of ductus deferens and epididymal tail. The solitary SP-positive fibers, occasionally encountered in the connective tissue of the tunica albuginea in the fallow deer and the pampiniform plexus of the red deer, can supplement the sensory functions of the C G R P - I R axons in both species. As has already been described for a number of mammals from different systematic groups, NPY is also the dominating neuropeptide in the cervine male gonad. NPY is often colocalized with D B H in the intramural arterial plexuses and some fibers reach also the walls of the veins in the spermatic cord. The arterial innervation pattern of the NPY-positive fibers very typically displays the variations that have been reported dependent upon the functional state of the seminiferous epithelium. It is widely accepted that activation of the postjunctional NPY-Y1 receptor elicits a vasoconstriction in arteries and veins, either directly or indirectly by potentiating the responses to noradrenaline (Newhouse and Hill 1997; Lang and Maron 1997; Barrios et al. 1998; Hokfelt et al. 1998; Kagstrom et al. 1998; Phillips et al. 1998; Kotecha 1998; Chu and Beilin 1998; Han et al. 1998) or by counterbalancing a VIP-induced vasodilation (Larsen et al. 1981; Polak and Bloom 1984; Kopp et al. 1997). In the testis of the deer, the cholinergic innervation is only sporadic and of minor functional importance. This finding corroborates the negative results obtained in the testes of other ungulates viz. bull (Wrobel and Abu-Ghali 1997), pig (Wrobel and Brandl 1998), donkey (Wrobel and Moustafa 2000) and camel (Saleh et al. 2002 a). Earlier reports, generally and regularly describing a rich cholinergic innervation of the mammalian testis (Risley and Skrepetos 1964; E1-Badawi and Schenk 1967; Bell and McLean 1973; Langford and Silver 1974) were based on suboptimal acetylcholinesterase techniques and are therefore not reliable. It must, however, be mentioned that the same advanced and sound methodology, used in this present investigation, demonstrated a remarkable cholinergic innervation of the medium-sized intralobular arteries in the feline testis (Wrobel and Gtirtler 2001). So, the question of a significant cholinergic testicular innervation has to be answered separately for each mammalian species. VIP-containing axons to the testis also display a remarkable species-specific distribution pattern, but generally seem to prefer the ISN and the caudal access route (Suburo et al. 2002). V1P-IR fibers are virtually absent in the testis of man, rat and donkey (Vaalasti et al. 1986; Rauchenwald et al. 1995; Wrobel and Moustafa 2000). In deer as in the camel (Saleh et al. 2002 a, b), only the ISN contains the V I P - I R fiber contingent. The majority of them end in mucosa and muscular coat of the deferent duct and epididymal tail, but some reach the testis proper as part of the caudal and mesorchial contributions. In fallow and roe deer, the main target region for VIP-IR fibers is the tunica albuginea. The testis of the red deer contains significally more VIP-positive axons which accompany the arteries into the testicular lobules where

they probably exert vasomotor functions, for VIP is known to be a potent vasodilator (Said 1984; Lissbrant et al. 1997). The small fraction of VIP-IR fibers that terminate in the red deer apparently independent of blood vessels near tubular walls and within Leydig cell groups could well be of a sensory nature, as has been concluded for such fibers in the cat testis (Suburo et al. 2002) or for non-vascular VIP-positive axons in other sites of the urogenital system (Dixon et al. 1994; R6sch et al. 1997).

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