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Departments of Medicine3 and of
Cell Biology and Physiology,4 University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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follicle-stimulating hormone, luteinizing hormone, spermatogenesis, testis, testosterone
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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While proliferation of undifferentiated spermatogonia continues throughout prepubertal development in the monkey [7], type B spermatogonia are generally not observed until the onset of puberty [7, 8]. Therefore, the differentiation of Ap spermatogonia to type B spermatogonia represents the crucial juncture in the initiation of spermatogenesis in the primate testis [4]. One to four generations of B spermatogonia are observed in the seminiferous epithelia of higher primates [4]. In rhesus monkey, four generations of type B spermatogonia (B1–4) have been identified [1]. The nuclei of these cells also have distinctive patterns of hematoxylin staining and are easily distinguished from the type A spermatogonia. The nuclei of each succeeding generation of type B spermatogonia have increasing amounts of heterochromatin [3, 4]. The sequential divisions of these premeiotic germ cells are chiefly responsible for production of the extraordinary numbers of spermatozoa that are characteristic of macaque spermatogenesis.
As in all mammalian species, spermatogenesis in higher primates is dependent on the gonadotropins, although the arrest of the process in the absence of LH and FSH is premeiotic in the latter species [9]. The purpose of the present study was to investigate, in the adult testis, whether self-renewal of Ap spermatogonia is gonadotropin dependent and, if so, the relative role of LH and FSH in this proliferative process. To this end, adult male rhesus monkeys were rendered hypogonadotropic by daily treatment with a GnRH-receptor (GnRH-R) antagonist, acyline, and later treated with testosterone (T) and FSH, either alone or in combination. In addition, this experimental design also provided an opportunity to examine the hormonal requirements for differentiation of Ap to B1 spermatogonia.
It should be noted that changes in circulating inhibin B concentrations and testicular inhibin ßB expression observed during the present study have already been published [10].
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Twenty adult male rhesus monkeys (Macaca mulatta; age, 4.5–8 yr; body weight, 5.5–11 kg; combined testicular volume, 17–24 ml) were used. The animals were maintained in accordance with the NIH Guidelines for the Care and Use of Laboratory Animals, and the experimental procedures were approved by the University of Pittsburgh Institutional Animal Care and Use Committee.
Surgical Procedures
The details of the surgical procedures, including implantation of i.v. catheters, testicular biopsies, and castration, have been published previously [10]. Postsurgically, all animals received a single i.m. injection of penicillin (300 000 U of Bicillin L-A; Wyeth Laboratories, Philadelphia, PA) and i.v. injections of a broad-spectrum antibiotic (100 mg of cefazolin sodium; Kefazol; Apothecon, Princeton, NJ) and an analgesic (1 mg/kg body weight of meperidine hydrochloride; Demoral; Elkins-Sinn, Cherry Hill, NJ) twice daily for 4 days. Implantation of T-filled or blank (B) Silastic capsules was performed under sedation with ketamine hydrochloride (100 mg i.m. and 50 mg supplements as required; Ketaset; Fort Dodge Laboratories, Fort Dodge, IA).
Access to Venous Circulation
The Silastic catheters were tunneled s.c. from the site of venous insertion (internal jugular and femoral vein) to the midscapular region, where they were exteriorized via a small cutaneous fistula. Animals with catheters were fitted with a nylon jacket attached via a flexible stainless-steel tether to a swivel device mounted on top of the cage, permitting continuous access to the venous circulation without tranquilization and with minimal restraint. One of the i.v. catheters was dedicated to FSH or vehicle (V) infusion, and the other was used for the withdrawal of blood samples. The routine care of the monkeys housed in the remote sampling laboratory has been described previously [11].
Collection of Blood
Before catheterization, blood samples were collected by femoral venipuncture following sedation with ketamine hydrochloride (100 mg i.m.). After catheterization, blood samples were drawn via the catheter into heparanized syringes. Serum and plasma were separated and stored at –20°C until required for assay.
Hormones and GnRH Antagonist
Recombinant human (rh) FSH was kindly provided by the National Hormone and Peptide Program (NIDDK, Harbor-UCLA Medical Center, Torrance, CA). The preparation of the rhFSH stock (200 IU/ml) and working solutions (2.2–2.9 IU/ml) have been described previously [10]. The T-filled or empty Silastic capsules (length, 5 cm; inner diameter, 3.35 mm; outer diameter, 4.65 mm; Dow Corning, Midland, MI) were prepared as described previously [12]. The GnRH-R antagonist, acyline (300 µg/ml; Bioqual, Rockville, MD), was prepared in 5% aqueous mannitol (AMVET Scientific Products, Yaphank, NY) and stored at 4°C [10].
Experimental Design
A schematic of the overall experimental design, which is described in three separate but interrelated phases, is presented in Figure 1.
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Phase 1: Establishing a chronic hypogonadotropic state Baseline values for circulating hormone concentrations were established for each of the 20 monkeys before the hypogonadotropic state was induced. For this purpose, one or more blood samples were collected at approximately 1000 h. The testicular volume of each animal was determined at this time [12]. A hypogonadotropic state was then induced and maintained by daily injections of the GnRH-R antagonist (acyline, 60 µg/kg). Following the initiation of GnRH-R antagonist treatment, the animals were sedated weekly to collect a blood sample and to measure testicular volume.
When the decline in testicular volume of an animal had plateaued for three consecutive weeks, between 15 and 26 wk after initiation of treatment with GnRH-R antagonist, a biopsy sample was obtained from one randomly chosen testis. The biopsy was fixed in Bouin fluid, embedded in paraffin, sectioned (thickness, 4 µm), and stained with periodic acid-Schiff (PAS)-hematoxylin for immediate histological examination as described previously [12]. Following confirmation of a full regression in spermatogenesis, the second phase was initiated.
Phase 2: T treatment After 20–33 wk of treatment with GnRH-R antagonist, and within 7 wk of the biopsy, when the reduction in testicular volume stabilized at approximately 10% of the pre-GnRH-R antagonist-treatment value, the monkeys were divided into two groups of 10 each. One group was implanted with T-filled capsules (20 capsules/monkey) to restore testicular content of this steroid, and the other group was implanted with empty or blank (B) capsules (20 capsules/monkey). The collection of blood samples and the measurement of testicular volume continued on a weekly basis.
Phase 3: rhFSH treatment After 11–44 wk of treatment with T or empty capsule implantation, all monkeys were fitted with indwelling i.v. catheters and housed in remote sampling cages. Five monkeys from each of the T- and non-T-treated groups were then stimulated for 12 days with an intermittent i.v. infusion of rhFSH (2 IU/kg per pulse, 2 ml as a 3-min infusion every 3 h). The remaining five monkeys from each of the T- and non-T-treated groups received an infusion of V. The four groups of five animals each are designated as follows: GnRH-R treatment only (B+V), T replacement alone (T+V), FSH replacement alone (B+FSH), and a combination of T and FSH replacement (T+FSH).
5-Bromo-2'-Deoxyuridine Injection and Collection of Testicular Tissue
On the last day of FSH or V infusion, all monkeys were bilaterally castrated. It should be noted that only four of the five monkeys in each group received a bolus i.v. injection of 5-bromo-2'-deoxyuridine (BrdU; 33 mg/kg; Sigma Chemical Co., St. Louis, MO), and these animals were castrated 3 h later [13]. Each testis was weighed and cut into several portions. Some of these were fixed in Bouin fluid overnight. Tissue from both testes also was frozen in liquid N2 and stored at –80°C for determination of testicular T content.
Gonadotropin Assays
Circulating concentrations of endogenous FSH and LH were determined using homologous (cynomolgous) RIA reagents supplied by the National Hormone and Peptide Program as described previously [14, 15]. The sensitivity of the FSH assay was 0.12 ng/ml, and the intra- and interassay coefficients of variation were less then 3% and 15%, respectively. The sensitivity of the LH assay was 0.11 ng/ml, and the intra- and interassay coefficients of variation were less than 4% and 9%, respectively.
Testosterone
Plasma T concentrations were measured using a commercially available solid-phase RIA kit (Total T [TKTT], Coat-A-Count; Diagnostic Products Corporation, LA). This assay also was used for measurement of testicular T concentration [16]. For the latter purpose, testicular tissue (74– 133 mg) was homogenized in a known amount of PBS-gel buffer, and the homogenate, containing a known amount of tritiated T (
2000 cpm) to determine recovery, was extracted with 5 ml of diethyl ether for 15 min with constant shaking. After evaporation of the ether, the extract was reconstituted in a known volume of PBS-gel buffer before assay. The mean sensitivity of the assay was 0.014 ng/ml, and the intra- and interassay coefficients of variation were less than 9% and 10%, respectively. The mean recovery from testicular tissue was 66% ± 22%.
Morphometric Analysis
Portions of the Bouin-fixed testicular tissue from each monkey were embedded in paraffin, and histological sections (thickness, 4 µm) were stained with PAS-hematoxylin. Two sections from each testis were selected randomly, and the germ and Sertoli cells in 300–500 cross sections of seminiferous tubules (ST), as defined by circular profiles, were counted. All cell counts were corrected by the method of Abercrombie [17] and the results expressed as cell number per cross section of seminiferous tubule (NS).
In addition, a further calculation was performed to determine the total number of Ad and Ap spermatogonia and Sertoli cells per testis [7]. First, the volume fractions (VV) of the interstitium and seminiferous tubules were determined by the point-counting method [18]. Briefly, a grid of intersecting lines of known area was superimposed over a section. The number of intersections on the grid (test points) overlying the tissue component of interest (Pn) were counted, and the ratio of these points to the total number (PT) was the volume fraction of that component (see Eqn. 1 below). A total of 4000 test points were examined on a randomly selected histological section. Second, the diameter (DS) of 25 cross sections of seminiferous tubules was measured with a calibrated ocular micrometer. Third, the seminiferous tubules were assumed to be cylindrical, and their lengths were estimated from the absolute volume (VS) and DS, as indicated in Equations 2 and 3, respectively. The number of the particular cell per cross section was then multiplied by total cord length to yield the number of this cell type per testis (Eqn. 4):
where L = length.
In the case of differentiated, type B spermatogonia, these four cell types were expressed only as the number per cross section of seminiferous tubule. This is because the length of the seminiferous tubule containing each generation must be determined from serial sections of the entire testis to calculate the total number of each generation of this cell type per testis. Figure 2 shows the typical morphology of each cell type.
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BrdU Labeling Index
The BrdU was visualized in two randomly selected histological sections (thickness, 5 µm) from each monkey using a previously described method [13]. Briefly, after washing the sections, they were incubated with mouse anti-BrdU antibody (Roche Diagnostic Corp., Indianapolis, IN). The sections were washed repeatedly and incubated with biotinylated horse anti-mouse antiserum, which was visualized with avidin-horseradish peroxidase complex (Vectastain ABC Elite Kit; Vector Laboratories, Inc., Burlingame, CA). The sections were counterstained with PAS. The number of labeled and unlabeled Ad, Ap, and B1–4 spermatogonia in cross sections of seminiferous tubules in which these cell types were identified was then determined. The labeling index of each cell type was calculated by dividing the number of labeled cells by the sum of both labeled and unlabeled cells (200–350 of each cell type per animal) and then expressed as a percentage. In the case of Ap spermatogonia, an additional labeling index was calculated using only those seminiferous tubule cross sections in which labeled Ap spermatogonia were observed. For those animals in which the stages of the seminiferous epithelium could be determined, the additional labeling index was derived from only stages VII–IX, when Ap spermatogonia are known to divide [1–3]. Finally, the percentage of cross sections having labeled Ap spermatogonia was calculated by dividing the number of cross sections with one or more labeled Ap spermatogonia by the total number of cross sections evaluated.
Statistical Analysis
A one-way ANOVA was used to compare the parameters of all four groups, and when indicated, the Student-Newman-Keuls multiple-range test was performed [19]. In the case of testicular T content, the data were not normally distributed and, therefore, were logarithmically transformed before performing ANOVA and, subsequently, the Student-Newman-Keuls multiple-range test. Significance was determined when P 
0.05. All data are expressed as the mean ± SD.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A detailed description of changes in hormone concentrations throughout the present study has been published previously [10]. In brief, treatment with GnRH-R antagonist resulted in a marked decrease of circulating endogenous gonadotropins to levels at or below the limit of detection, whereas T levels were reminiscent of those in castrated monkeys (i.e., 0.2 ng/ml) [20]. In the monkeys implanted with T-filled Silastic capsules (T+V and T+FSH), circulating concentrations of this steroid were maintained at 15 ng/ml for the duration of the present study. In the groups receiving rhFSH infusion, the peak concentrations of this gonadotropin in the circulation ranged from 19–30 IU/L.
Testicular Parameters
The mean testicular weight of the monkeys treated with T+FSH was significantly greater than those of the other three groups of monkey (Table 1). The mean testicular weights of the monkeys treated with FSH or T alone (B+FSH and T+V) were not significantly different from each other or from those of animals treated with GnRH-R antagonist only (B+V). The mean seminiferous tubule diameter was significantly larger in the animals receiving T+FSH (Table 1). The mean total lengths of the seminiferous tubules were not different among the four groups of monkeys, and as expected, the mean number of Sertoli cells per testis was statistically indistinguishable (Table 1).
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The mean testicular T contents of the monkeys treated with T and FSH, either alone or in combination, were not significantly different from the mean testicular T content of a group of five normal monkeys (Table 2). However, the T contents of the animals receiving only the GnRH-R antagonist (B+V) were significantly different from those of the other treated groups (T+V, B+FSH, and T+FSH) and those of the normal monkeys.
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Seminiferous Epithelium
A representative photomicrograph from one monkey in each of the four groups is shown in Figure 3. Within each group, the histological appearance of the seminiferous epithelium of the individuals was similar. The seminiferous epithelia of the monkeys treated with GnRH-R antagonist alone (B+V) comprised only Ad spermatogonia, Ap spermatogonia, and Sertoli cells (Fig. 3A). The epithelia of the monkeys receiving T, either alone or in combination with FSH, were composed of all germ cell types, including elongated spermatids, and these were frequently arranged in identifiable cellular associations representing stages of the cycle of the seminiferous epithelium (Fig. 3, C and D). The seminiferous epithelia of the monkeys implanted with empty capsules and receiving FSH for only 12 days comprised, in addition to the undifferentiated spermatogonia and Sertoli cells, all four generations (B1–4) of differentiated spermatogonia and meiotic germ cells as mature as zygotene to early pachytene spermatocytes (Fig. 3B). Within any given cross section of a seminiferous tubule from FSH-treated animals, usually only one generation, but on occasion two successive generations, of B spermatogonia were observed.
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Morphometry of the Seminiferous Epithelium
Examination of the total number of Ap spermatogonia per testis and the labeling index of this cell type provided no evidence to indicate that proliferation of Ap spermatogonia was stimulated by T and FSH, either alone or in combination (Fig. 4). The mean numbers of Ap spermatogonia per tubule cross section or per testis of the four treatment groups were not significantly different (Fig. 4). In groups receiving T, labeled Ap spermatogonia were observed in the expected stages (VII–IX), and using all cross sections exhibiting these stages, the labeling index of Ap spermatogonia was 55.4% ± 11.5% and 48.0% ± 14.5% in T+V and T+FSH, respectively. In the two groups in which stages were not observed (B+V and B+FSH), the labeling index was derived from sections in which S-phase labeling of this cell type was observed. The labeling indices were 56.5% ± 2.7% and 50.7% ± 12.0%, respectively, and no statistical differences between GnRH-R antagonist (B+V) and groups treated with FSH and T, either alone or in combination (Fig. 4). In the two groups in which stages were not present, the percentage of cross sections showing BrdU-labeled Ap spermatogonia was 4.8% ± 2.2% and 7.2% ± 2.2% in B+V and B+FSH, respectively, compared to values of 5.8% ± 5.2% and 10.8% ± 5.5% in T+V and T+FSH, respectively, the groups in which stages were present. These percentages were not significantly different.
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The labeling of Ap spermatogonia in randomly selected cross sections, regardless of stages (the two T-treated groups), ranged from 4% (B+V) to 12% (T+FSH). Again, this parameter did not show any statistical differences among the four groups.
In the case of Ad spermatogonia, quantitative analysis revealed that the mean number of these cells ranged from 0.43 ± 0.19 to 0.59 ± 0.24 per cross section, or 70 ± 14 x 106 to 97 ± 21 x 106 per testis, and were not significantly different among the four groups. The labeling index of Ad spermatogonia in randomly selected cross sections was low and independent of treatment (B+V, 1.3% ± 0.5%; T+V, 1.2% ± 0.8%; B+FSH, 2.3% ± 2.0%; T+FSH, 0.7% ± 0.8%).
The mean number of each generation of differentiated type B spermatogonia per cross section is shown in Figure 5. Type B spermatogonia were not observed in the testes of the monkeys receiving only GnRH-R antagonist (B+V). In contrast, all four generations of type B spermatogonia were observed in the remaining three groups of monkeys (Fig. 5). In the monkeys treated with T and FSH in combination, the mean number of each generation of differentiated spermatogonia was significantly greater than those in the other three groups (Fig. 5). In the monkeys treated with either T or FSH alone, no significant differences were observed in mean number of B1, B2, B3, and B4 spermatogonia.
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The percentage of differentiated spermatogonia of each generation incorporating BrdU is presented only for those animals in which type B spermatogonia were observed (Table 3). The percentages of labeled B1, B2, B3, and B4 spermatogonia ranged between 25% and 44% and were not statistically different among the three groups of monkeys in which they were observed. The proportion of a particular generation of type B spermatogonia incorporating BrdU was the same irrespective of the treatment, although the mean number of the particular differentiated spermatogonium was different with respect to treatment.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The primary purpose of the present study was to determine the relative role of LH and FSH in the self-renewal and proliferation of Ap spermatogonia in this representative higher primate. In this regard, the finding that the Ap spermatogonia in the chronically hypogonadotropic group robustly incorporated BrdU was particularly striking. Indeed, mitotic activity in Ap spermatogonia in the hypogonadotropic group was not significantly different from those of the groups treated with T and FSH, either alone or in combination. This result is consistent with our recent observation that incorporation of BrdU by Ap spermatogonia during juvenile development in the monkey, a naturally hypogonadotropic state in these species, is similar to that during infancy, when LH and FSH secretion is elevated (unpublished observation). Taken together, these findings lead to the conclusion that in the rhesus monkey, a marked level of mitosis of Ap spermatogonia continues in the absence of a gonadotropin drive. A similar situation may pertain in New World monkeys, because treatment with a GnRH-R antagonist during infancy had only a minor impact on the total number of spermatogonia [21]. In the rat, a similar inference may be made for type A spermatogonia, because germ cells as mature as step 8 spermatids are observed in the testes of adult rats rendered chronically hypogonadotropic [22, 23]. Therefore, a major component of undifferentiated spermatogonial proliferation in mammalian species, in general, may occur in a constitutive manner in the absence of stimulation by the pituitary gonadotropins. This constitutive proliferation is, presumably, amplified by LH and/or FSH, because the number of Ap spermatogonia in the adult testis of the macaque is reduced following the induction of a chronically hypogonadotropic state. In the present study, the total number of this cell type in the hypogonadotropic group was approximately twofold lower than that previously observed by us in the testes of normal adult rhesus monkeys [7]. A decrease in Ap spermatogonial number also has been noted previously by others following surgical removal of the pituitary or after treatment with either a GnRH-R antagonist or a contraceptive dose of T [24–27].
The question, therefore, becomes why the hormonal replacements employed in the present study did not increase the number of Ap spermatogonia. The failure to amplify Ap spermatogonial number in the hypogonadotropic group is unlikely to result from inadequate T replacement, because the testicular T contents of the T-treated monkeys were indistinguishable from those of normal intact monkeys, as in the present study and as reported previously [16]. Because in the crab-eating monkey initiation of T replacement immediately after hypophysectomy maintained the number of Ap spermatogonia [24], the failure of T replacement to restore Ap spermatogonial number may be related to the protracted 6- to 8-mo period of hypogonadotropism before T replacement was initiated.
Similarly, previous studies of the crab-eating monkey have shown that initiation of FSH treatment concomitant with that of a GnRH-R antagonist also may prevent the depletion of Ap spermatogonia [25]. Moreover, in the latter experimental paradigm, delayed administration of an 8-wk regimen of FSH treatment resulted in a partial restoration of the number of Ap spermatogonia [25]. We are reluctant to conclude, however, that the short duration of FSH treatment (12 days) used in the present experiment can explain the failure of this gonadotropin to increase the Ap spermatogonia population, because differentiation of Ap spermatogonia and survival of their progeny, B1 spermatogonia, were observed.
Therefore, we are left with the protracted period of hypogonadotropism before initiation of hormone replacement as the most parsimonious explanation for the failure of the regimens of T and/or FSH treatment to increase the number of Ap spermatogonia in the present study. Perhaps the prolonged hypogonadotropism produced in the present experiment resulted in a change of testicular architecture, leading to a decreased availability of the stem cell niches [28] that was not restored by the hormonal replacement.
The present study also provided an opportunity to examine the relative role of the gonadotropins in governing the differentiation of Ap spermatogonia into B1 spermatogonia. The absence of type B spermatogonia in the monkeys treated with GnRH-R antagonist alone confirms the classic observation of Smith [22] that in the primate, differentiation of Ap to type B spermatogonia is gonadotropin dependent. Our finding that replacement with T alone led to production of type B spermatogonia similarly confirms those of earlier studies [24, 29, 30]. In adult, hypophysectomized crab-eating monkeys, differentiated spermatogonia were observed after 8 or 13 wk of T replacement [24]. In addition, treatment of juvenile rhesus monkeys with either T [30] or LH [29] resulted in production of differentiated spermatogonia. In the former case, the germ cells were not enumerated, but the presence of germ cells more mature than type B spermatogonia supports the inference that the less mature, differentiated spermatogonia also were stimulated.
In addition, the present finding that FSH alone also was able to support the differentiation of Ap spermatogonia substantiates the results of other studies involving the adult crab-eating macaque [25] and is consistent with the observation that FSH treatment of the juvenile rhesus monkey resulted in the production of differentiated type B spermatogonia [29]. We conclude from the foregoing considerations, therefore, that FSH alone is sufficient for this key step of differentiation in primate spermatogenesis. Because FSH treatment alone was associated with an increased testicular T content, the possibility that activation of androgen receptor may be involved in mediating this action of FSH cannot be excluded.
In the present study, not only did either T or FSH permit the differentiation of Ap spermatogonia into B1 spermatogonia, these regimens also allowed the production of three subsequent generations of differentiated spermatogonia (i.e., B2, B3, and B4). Moreover, T and FSH were equally effective in stimulating the number of each of these cell types. The combination of T and FSH resulted in augmentation of the effects of either FSH or T alone on the number of each generation of differentiated spermatogonia. Indeed, treatment with T and FSH together produced yields of differentiated spermatogonia that closely approximated the theoretical yields.
The finding that Ad spermatogonia incorporated BrdU at a labeling index of less than 2.5% is entirely consistent with dogma that this cell type is a reserve stem cell [1–3] that is usually quiescent and becomes active only when the seminiferous epithelium is catastrophically damaged [4]. It should be noted that the number of Ad spermatogonia, as in the case of Ap spermatogonia, in the chronically hypogonadotropic animals also was approximately twofold lower than that in intact animals [7, 16].
Finally, the high testicular content of T in association with low circulating levels of the steroid in the animals receiving FSH alone merits comment. The most likely explanation for this finding is that FSH results in retention of T within the testis. In this regard, sex hormone-binding globulin is found in the primate testis and binds T with high affinity [31]. By analogy to the rodent, FSH may stimulate the primate Sertoli cell to produce this androgen-binding protein by a mechanism analogous to that responsible for FSH stimulation of androgen-binding protein in the rat [31]. The reasons for the high testicular T content in the FSH-treated group requires further study, but this interesting finding does not detract from the major conclusion of the present study that the proliferation of Ap spermatogonia is largely constitutive and independent of gonadotropin stimulation.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Gary R. Marshall, University of Pittsburgh, E1140 BST, 200 Lothrop Street, Pittsburgh, PA 15261. FAX: 412 692 4155; marshg{at}pitt.edu
Received: 15 December 2004.
First decision: 6 January 2005.
Accepted: 1 March 2005.
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REFERENCES |
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