1 Introduction

It was found in our previous study that a progressive degeneration of seminiferous tubules could be induced by testicular capsulotomy^[1]. However, the underlying mechanism is not clear. Although it is well-known that gonadotrophin and testosterone are fundamental for initiating, maintaining and regulating spermatogenesis^[2,3], we are uncertain whether the spermatogenetic disruption in capsulotomized testis is causally related to the changes in these hormones. On the other hand, it is possible that the pathological alterations of the testis induced by testicular capsulotomy may affect testosterone production, and exert a feedback influence on the gonadotrophins.

2 Materials and methods

2.1 Experimental animals and testicular capsulotomy

Mature (60 days of age) Sprague-Dawley male rats were obtained from the Laboratory Animal Unit of The University of Hong Kong and divided into control, sham-operated and capsulotomized groups of 6-8 rats each. Surgical intervention was performed in the Minimal Disease Operation Theatre of the University. The animals were anaesthetized intraperitoneally with sodium pentobarbitone (Sigma, USA) at a priming dose of 60 mg/kg and a maintenance dose of 10 mg/kgh. Testicular capsulotomy was carried out as previously described^[1]. Briefly, with the aid of a dissecting microscope (Wild M60, Switzerland), the two outer layers of the capsule, i.e., tunica vaginalis and tunica albuginea,  were carefully incised starting half-way down the rostral half of the testis along the two lateral borders down to the middle of the caudal half of the testis.

2.2 Testosterone determination

Testosterone was measured by radioimmunoassay (RIA) according to the method of Nankin and Troen^[4]. The kits were purchased from Amersham (UK).

The animals were anaesthetized intraperitoneally with sodium pentobarbitone (60 mg/kg, Sigma, USA) and then heparinized (1250 U heparin, Sigma, USA) via the femoral vein. Testis was exposed through a scrotal incision. The band of tissue joining the head of the epididymis and the testis was severed to reveal the testicular venous plexus. A vein just distal to this plexus was punctured with a 25-gauge needle and blood was collected into a heparinized tube^[5,6]. The samples were stored overnight at 4. Plasma was separated by centrifugation at 1500g for 30 min at -4, and then stored at -20 until determination. Testosterone levels were measured at 10-day intervals post-operation for a period of 60 days.

The recovery of the added tritium labeled-testosterone was in the range 80%-86%. Samples from any single experiment were always assayed in a single assay and the intra- and inter-assay coefficients of variation were <10%.

2.3 LH and FSH determination

LH and FSH were measured by double antibody radioimmunoassay as previously described^[7]. A catheter was inserted into one jugular vein from which blood samples were drawn.The RIA kits for rLH and rFSH were purchased from Amersham (UK).

Samples from any single experiment were always assayed together in a single assay and the intra- and inter-assay coefficients of variation were always <9%.

2.4 Statistical analysis

3 Results

3.1 Testosterones

The testosterone levels in testicular vein showed the same pattern of change in the normal and sham-operated rats. In both groups the concentrations of testosterone slightly increased at the age of Day 80 and then decreased significantly (P<0.05) at the age of Day 90. From 90 to 120 days of age, the testosterone values remained relatively constant.  The testosterone levels in testicular vein of the sham-operated animals were not significantly different from those of the normal rats during the period of study (Figure 1).

Figure 1. Testosterone level in testicular vein of normal and sham-operated rats at 10-day intervals. Filled histograms, normal rats. Open histograms, sham-operated rats. meanSEM, from a group of 8 rats.

For the capsulotomized rats, the testosterone levels in the testicular vein were increased  at 20 days post-operation in comparison to those of capsulotomized rats at 10 days post-operation. However, from 30 to 50 days after capsulotomy, the testosterone concentrations were significantly (P<0.05) reduced to 95-83% of those at 10 days post-capsulotomy. At the last observation point (60 days after capsulotomy), the testosterone concentration were significantly (P<0.05) reduced further to approximately 56% of those at 10 days post-capsulotomy.

The testosterone levels of the capsulotomized rats were significantly different from those of the sham-operated control from 30 days onwards (Figure 2). At 30 days after capsulotomy, the testosterone levels were reduced to 76% of the corresponding control value (P<0.05). By 60 days it had dropped to only 49% of the corresponding control (P<0.05).

Figure 2.  Testosterone level in testicular vein of sham-operated and testicular capsulotomized rats at various intervals post-operation. Filled histograms, sham-operated rats. Open histograms, capsulotomized rats. Values are  meanSEM, from a group of 8 rats. ^bP<0.05, compared with corresponding sham-operated controls.

3.2 LH and FSH

With increase in age, the plasma concentrations of LH and FSH rose significantly in both groups of sham-operated control rats and capsulotomized rats, and remained elevated during the period of study.  However, more remarkable time-dependent increase of these hormones was observed in capsulotomized rats from 20 to 60 days post-operation in comparison to sham-operated rats.

For the sham-operated rats, plasma concentrations of LH were significantly (P<0.05) increased by 12% at 20 days, 26% at 40 days, and 37% at 60 days respectively in comparison to those of sham-operated rats at 10 days. For the capsulotomized rats, however, they were significantly (P<0.05) increased by 31% at 20 days, 69% at 40 days, and 83% at 60 days respectively when compared with those of capsulomized rats at 10 days.

Plasma concentrations of FSH in the sham-operated rats were significantly (P<0.05) increased by 7% at 20 days, 40% at 40 days, and 43% at 60 days respectively in comparison to those of sham-operated rats at 10 days. However, for the capsulotomized rats, they were significantly (P<0.05) increased by 32% at 20 days, 98% at 40 days, and 89% at 60 days respectively when compared with those of capsulotomized rats at 10 days.   

For the capsulotomized rats, plasma concentrations of LH and FSH at 10 days post-operation were 35 and 247 ng/mL, respectively, which were not significantly different from those of their control groups. However, both LH and FSH levels were significantly elevated 20 days post-capsulotomy when compared with the control rats; LH levels increased from 40 ng/mL to 46 ng/mL (P<0.05) and FSH from 277 ng/mL to 327 ng/mL (P<0.05). These levels were elevated further by day 40 post-capsulotomy; plasma LH increased from the control value of 45 ng/mL to 59 ng/mL (P<0.05) and FSH from 360 ng/mL to 490 ng/mL (P<0.05). By 60 days post-capsulotomy, LH and FSH levels remained elevated at 64 and 467 ng/mL, respectively. (Figure 3 and 4).

Figure 3.  Peripheral plasma level of LH in sham-operated and testicular capsulotomized rats at various intervals post-operation. Filled histograms, sham- operated rats. Open histograms, capsulotomized rats. Values are  meanSEM, n=6, ^bP<0.05,  compared with corresponding sham-operated controls.  
Figure 4.  Peripheral plasma level of FSH in sham-operated and testicular capsulotomized rats at various intervals post-operation. Filled histograms, sham-operated rats. Open histograms, capsulotomized rats. Values are  meanSEM, n=6, ^bP<0.05,  compared with corresponding sham-operated controls.

4 Discussion

We observed a progressive reduction in testosterone level in the testicular venous blood in rats after capsulotomy. Significant changes in testosterone level occurred around 30 days after treatment. Since considerable morphological changes were observed at 5-10 days after capsulotomy, i.e., prior to the testosterone change^[1], it seems that the seminiferous tubular degeneration after testicular capsulotomy was not caused by the reduction in testosterone.

It is well known that the spermatogenic process is extremely sensitive to changes in the blood supply to the testis. Disruption of the spermatogenesis can result rapidly from ischemia^[8], and the production of testosterone by Leydig cells can also be influenced by altered blood flow^[9]. Testicular capsulotomy will possibly cause a change in the mechanical environment of the testis and this in turn may influence the tone of the blood vessels and the amount of blood supply to the testis. Hence, the blood supply to the testis was monitored. For rats which were sham-operated or had their testes capsulotomized at the age of Day 60, the blood flow of testicular artery was measured directly by ultrasonic flowmeter at 10-day intervals throughout post-operation period. However, there were no significant changes in the testicular blood flow of both groups of rats. The result indicated that the alteration in the concentrations of testosterone in testicular venous blood of capsulotomized rats resulted from other mechanism(s).

In recent years, there has been accumulating evidence indicating that the action of LH on the Leydig cells is dependent on the local environment created through interactions between the seminiferous tubules and the interstitial cells. Paracrine factors are believed to be the major means for the communication involved^[10]. Hence, a significant reduction in testosterone level occurring after seminiferous tubular degeneration in capsulotomized testis is most likely caused by changes in local paracrine factors. Recently, the paracrine control of Leydig cells by factors released from the seminiferous tubules has been the subject of multifarious investigations^[11]. Both inhibitory (activin and TGF-) and stimulatory (inhibin and IGF-1) factors have been identified in in vitro preparations^[12-16]. Wu and Murono^[17] have found an as yet unidentified testicular growth factor(s) which stimulates proliferation but inhibits steroidogenesis of the rat Leydig cells. It is possible that the degenerating seminiferous tubules may release such kind of factor(s) causing inhibition of testosterone production.

Kerr et al^[18] attributed the decrease in testosterone in these situations to a rapid metabolism of the hormone by the damaged seminiferous tubules. Some workers have shown that Sertoli cells under the influence of FSH can metabolize testosterone to oestradiol^[19,20]. Hence, it is very likely that in the capsulotomized testis, the metabolic status of the Sertoli cell is changed under  high FSH concentrations, that favours the conversion of testosterone to oestradiol.

In rats with testicular capsulotomy, although testosterone within the testicular vein was found to be decreased, its level was still within the range of 42.5-49.7% of the control value (Figure 2). Such a level of testosterone is adequate for maintaining normal spermatogenesis^[3]. Hence, it is doubtful that the reduction of testosterone can have  any directly adverse effects on spermatogenesis in the capsulotomized testis. However, it is possible that the reduction in testosterone may facilitate the degenerative changes of the seminiferous tubules induced by other factors.

Although our experimental results suggest that the spermatogenic disruption was not directly the result of a reduction in testosterone production after capsulotomy, we cannot rule out the possibility that other functions of the testis, such as seminiferous tubular contraction and tubular fluid secretion may be influenced by the lowered testosterone level.

In vitro studies have demonstrated that high concentrations of testosterone induce contraction of the seminiferous tubules, whereas lower concentrations cause relaxation^[21]. Thus, the low testosterone level after testicular capsulotomy may cause seminiferous tubular relaxation. If contraction of the seminiferous tubules does play a role in sperm transport, its inhibition under low testosterone level may result in a slowing of sperm transport within the testis. The secretion of seminiferous tubular fluid by the Sertoli cells is considered by some workers to be involved in sperm transport and more importantly in the maintenance of the microenviroment for spermatogenesis^[22,23]. A number of studies in adult rats have shown that the production of seminiferous tubular fluid is under the control of testosterone^[24,25]. In the capsulotomized testis, the reduced level of testosterone may decrease tubular fluid secretion, and this will not only slow down sperm movement leading to sperm congestion inside the testis but also will result in a microenvironment unfavorable for spermatogenesis.

In testes of capsulotomized rats, we found a progressive increase in the LH and FSH levels from 20 days post-operation. The increase in LH level is most likely a result of negative feedback regulation of the lowered testosterone level, while the increase in FSH secretion may be principally due to a feedback signal from the damaged seminiferous tubules. It was reported that in many states of spermatogenic damage, the Sertoli cells are found to produce less inhibin but more activin, a FSH-releasing factor^[26,27]. Under such a situation, FSH released from the pituitary is increased significantly.

Acknowledgements

References

[1] Qin DN, Lung MA. Studies on the relationship between testicular capsule and sperm  transport in the rat testis. Asian J Androl  2000; 2: 191-8.
[2] Chemes HE, Dym M, Raj HGM. The role of gonadotropins and testosterone on initiation of spermatogenesis in the immature rat. Biol Reprod 1979; 27: 183-8.
[3] Sharp RM, Donachie K, Cooper I. Re-evaluation of the intratesticular level of testosterone required for quantitative maintenance of spermatogenesis in the rat. J Endocrinol 1988; 117: 19-26.
[4] Nankin HR, Troen P. Evaluation of testosterone in the male. In : Hafez ESE, editor. Techniques of human andrology.  Holland: Elsevier Biochemical Press; 1977, p 239-50. 
[5] Galil KAA, Setchell BP. Effect of local heating of the testes on the concentration of testosterone in jugular and testicular venous blood of rats and on testosterone production in vitro. Int J Androl 1987; 11: 61-72.
[6] Maddocks S, Setchell BP. The composition of extracellular interstitial fluid collected with a push-pull cannule for the testis of adult rats. J Physiol 1988; 407: 363-72.
[7] Lee VWK, de Krester DM, Hudson B, Wang C. Variation in serum FSH, LH, and testosterone levels in male rats from birth to sexual maturity. J Reprod Fertil 1975;  42: 121-6.
[8] Steinberger E, Tjioe DY. Spermatogenesis in rat testis after experimental ischemia. Fertil Steril  1969; 20: 639-49.
[9] Damber JE, Janson PO. The effect of LH, adrenaline and noradrenalin on testicular blood flow and plasma testosterone concentrations in anaesthetized rat. Acta Endocrinol Copenh 1978; 88: 390-6.
[10] Saez JM, Perrard-Sapari MN, Chatelain PG, Tabone E, Rivarola MA. Paracrine regulation of testicular function. J Steroid Biochem 1987; 27: 317-29.
[11] Bergh H. Local differences in Leydig cell morphology in the adult rat testis: evidence for a local control of Leydig cell by adjacent seminiferous tubules. Int J Androl 1982; 52: 325-30.
[12] Hsueh AJM, Dahl KD, Vaughan J. Heterodimers and homodiners of inhibin subunits have different paracrine actions in the production of luteinizing hormone stimulated androgen biosynthesis. Proc Natl Acad Sci USA  1987; 84: 5082-6.
[13] Kasson BG, Hsueh AJM. Insulin-like growth factor I augments gonadotropin stimulated androgen biosynthesis by cultured rat testicular cells. Mol Cell Endocrinol  1987; 52: 27-34.
[14] Lin T, Blaisdell J, Haskell JF. Transforming growth factor inhibits Leydig cell steroidogenesis in primary culture. Biochem Biophys Res Commun 1987; 146: 387-94.
[15] Rommerts FFG, Hoogerbrugge JW, Van der Molen HJ. Stimulation of steroid production in isolated rat Leydig cells by unknown factors in testicular fluid differs from the effects of LH and LH-releasing hormone. J  Endocrinol  1986; 109: 111-7.
[16] Sharpe RM, Cooper I. Intratesticular secretion of a factor(s) with major stimulating   effects on Leydig cell testosterone secretion in vitro. Mol Cell Endocrinol 1984; 37: 159-68.
[17] Wu N, Murono EP. A Sertoli cell secreted paracrine factor(s) stimulate proliferation and inhibit steroidogenesis of rat Leydig cells. Mol Cell Endocrinol 1994; 106: 99-109. 
[18] Kerr JB, Rich KA, de Kretser DM. Alterations of the fine structure and androgen secretion of the interstitial cells in the experimentally cryptorchid rat testis. Biol Reprod 1979; 20: 409-22.
[19] Armstrong DT, Moon YS, Tritz IB, Dorrington JH. Synthesis of estradiol-17beta by Sertoli cells in culture: stimulation by FSH and dibutyryl cyclic AMP. Curr Top Mol Endocrinol  1975; 2: 85-96.
[20] Dorrington JH, Armstrong DT. Follicle-stimulating hormone stimulates estradiol-17beta synthesis in the cultured Sertoli cells. Proc Nat Acad Sci USA 1975; 72: 2677-81.
[21] Yomamoto M, Nagai T, Takaba H, Hashimoto J, Miyake K. In vitro contractility of human seminiferous tubule in response to testosterone, dihydrotestosterone and estradiol. Urol Res 1989; 17: 265-8.
[22] Setchell BP.  The movement of fluids and substances in the testis. Aust J Biol Sci  1987; 39: 193-207.
[23] Waites GMH, Gladwell R. Physiological significance of fluid secretion in the testis and blood-testis barrier. Physiol Rev 1982; 62: 624-71.
[24] Au CL, Iry DC, Robertson DM, de Kretser DM. Effect of testosterone on testicular inhibin and fluid production in intact and hypophysectomized adult rat. J Reprod Fertil 1986; 76: 257-66.
[25] O' Leary PC, Jackson AE, Irby DC, de Kretser DM. Effect of ethane dimethane sulphonate (EDS) on seminiferous tubule function in rats.  Int J Androl 1987; 10: 625-34.
[26] Gonzales GF, Risbriger GP, de Krester DM. In vivo and in vitro production of inhibin by cryptorchid testes from adult rats. Endocrinology 1989; 124: 1661-8.
[27] Rivier C, Menuier H, Robert V, Vale W. Possible involvement of inhibin in alterted FSH secretion during dissociated LH and FSH release: unilateral castration and experimental cryptorchidism. Biol Reprod 1989; 41: 967-81.
[28]  Sharpe RM. Regulation of spermatogenesis. In: Knobil E,  Neill JD, editors. The   physiology of reproduction. New York: Raven Press, Ltd; 1994. p1363-434.