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Biology of Reproduction 60, 14-22 (1999)
©Copyright 1999 Society for the Study of Reproduction, Inc.

Stage-Dependent Accumulation of Cadmium and Induction of Metallothionein-Like Binding Activity in the Testis of the Dogfish Shark, Squalus acanthias1

M. Betkaa, and G.V. Callard2,a

a Department of Biology, Boston University, Boston, Massachusetts 02215


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cadmium (Cd) is an established spermatotoxicant. Using the shark (Squalus acanthias) testis model, we investigated stage-related patterns of intratesticular Cd accumulation and effect. After a single injection of 109CdCl2, tracer was rapidly eliminated from plasma but accumulated and was retained in testis for at least 7 days. Intratesticular 109Cd was stage dependent, resulting in a 3- to 5-fold gradient: germinal zone (GZ) > premeiotic (PrM) > meiotic (M) > postmeiotic (PoM) stages. When measured as tissue:plasma ratios, the Cd-binding mechanism in GZ (71:1) was similar to that in liver (87:1) but lower than in kidney (381:1). The same intratesticular gradient was seen in untreated controls when tissue Cd levels were measured by atomic absorption spectroscopy, implying environmental exposure. A single CdCl2 injection (5 mg/kg i.v.) elevated testicular Cd > 160-fold in all stages but did not alter the direction or magnitude of the gradient. Intratesticular distribution of metallothionein-like Cd-binding protein was stage dependent (PrM = PoM > GZ = M), but the pattern differed from the Cd gradient. This binding component was Cd inducible in all but M stages, but induction did not alter the stage-dependent pattern of binding activity or Cd accumulation. Analysis of tissue subfractions after in vivo tracer injection indicated that the binding mechanism responsible for the intratesticular gradient is mainly cytosolic, but that a second less abundant component is associated with the nucleus. The functional significance of preferential Cd accumulation in GZ and PrM stages of spermatogenesis remains to be determined.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Epidemiological research and studies in wildlife have reported an increase in male reproductive abnormalities, including increases in testicular cancer and declining sperm counts [1–3]. These reports have stimulated a debate as to the role of environmental chemicals, but clear cause-and-effect relationships remain to be established. What has hampered attempts to elucidate factors and mechanisms that control spermatogenesis is the complex organization of the testis of man and conventional laboratory and domestic animals. In these species, the number of different somatic elements that support spermatogenesis (Leydig, Sertoli, and peritubular myoid cells), and the arrangement and spacing between successive germ cell stages, make it technically difficult to analyze and experimentally manipulate a single discrete stage without disrupting germ cell clones, somatic elements, and the privileged germinal compartment [4, 5]. For the same reason, attempts to develop an in vitro spermatogenesis system for routine chemical testing and mechanistic studies have had only limited success [5]. Future progress in understanding human health effects, as well as the impact of environmental factors on normal spermatogenic efficiency and survival of animal species, has been linked to the acquisition of new experimental models and test systems [6].

The strategy of seeking simpler animal models is justified on the basis that spermatogenesis has been fundamentally conserved throughout the vertebrates, especially at the cellular and molecular levels [4, 7, 8]. Where phyletic and species differences do occur is in the organization of the testis. Studies in this laboratory have shown that the spiny dogfish shark (Squalus acanthias) is an ideal model for the step-wise analysis of spermatogenesis [9, 10]. Compared to other vertebrates, Squalus and related species have a unique combination of features: a) a "cystic" mode of spermatogenesis, in which a single germinal clone and a second clonal population of stage-synchronized Sertoli cells form a follicle-like unit (spermatocyst); b) the arrangement of spermatocysts in maturational order across the diameter of the testis, resulting in a readily visible zonation in testicular cross sections; and c) reliance mainly or exclusively on Sertoli cells for somatic cell support of spermatogenesis, differentiated Leydig cells being absent at this phyletic level (see Fig. 1). The present study was designed to test the utility of the shark testis model for stage-by-stage analysis of toxicant accumulation and effect, using cadmium (Cd) as an illustrative toxicant.



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  		FIG. 1. Organization of spermatogenesis and stages of spermatocyst development in shark testis. The diagrammatic cross section shows the simple diametric progression of spermatogenesis and demarcation of dissected tissues: GZ, PrM, M, PoM (see text). ZD, with a band of apoptotic cysts reflecting the preceding winter's period of spermatogenic inactivity, appears between PrM and M zones in May/June and was excluded from analyses except where indicated. The EO, a lymphomyeloid tissue, encapsulates the testes at the mature pole. Fluorescence confocal images show spermatocysts representative of staged tissues after staining with AO and their approximate position in the developmental progression: A) GZ with a nest of primitive cysts containing gonocytes (Go); B) PrM zone with cyst containing immature spermatogonia (Sg); C) PrM zone with cyst containing mature Sg; D) M zone with cyst containing spermatocytes (Sc); and E) segment of PoM cyst containing bundles of mature spermatids (St), with heads pointed peripherally and tails projecting into the lumen. Sertoli cell (SC) nuclei are not readily evident in D but are approximately equivalent in number to germ cells in A and B, located adluminally at the stage of maturation shown in C, and are adjacent to the basement membrane in E.

Cd is rare in nature and has no known biological utility [11]. Although Cd seldom reaches toxic concentrations in the environment, levels have progressively increased as a consequence of industrial pollution attributed to electroplating, stabilizers, pigments, plastics, semiconductors, and batteries [12, 13]. Moreover, the long half-life (10–30 yr in human kidney) and bioaccumulation and biomagnification of Cd in plants and animals increase the potential for human exposure and effect [11]. The rodent testis is exquisitely sensitive to Cd toxicity [14], and permanent or temporary sterility is induced at Cd doses below those that have adverse effects on the vasculature [15, 16]. The primary goal of the present study was to define the intratesticular distribution of radiolabeled and radioinert Cd as a function of developmental stage. Because metallothionein (MT) is a Cd-inducible, Cd-binding component [17], we used a standard Cd-binding assay to measure MT-like protein as a possible marker of tissue Cd accumulation and effect.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals

Mature male spiny dogfish sharks (Squalus acanthias; 1.06–2.18 kg; 36- to 49-cm snout-vent length) were collected from the Gulf of Maine through the facilities of the Mount Desert Island Biological Laboratory (Salsbury Cove, ME) during the period of spermatogenic activity (June–October) and were kept in running-seawater tanks no longer than 2 wk. Animals were killed by double pithing via the olfactory canal. Protocols for the care and use of animals were approved by the Mount Desert Island Biological Laboratory Institutional Animal Care and Use Committee.

Tissue 109Cd Concentrations and Organ Distribution

Animals (3 per treatment group) were injected with 109CdCl2 (specific activity 3.19 mCi/mg; New England Nuclear, Boston, MA; 2.84 µCi/0.3 ml water per animal) via the caudal vein. The estimated dose (2 µCi/kg) was based on a mean projected body weight of 1.4 kg. To determine blood and plasma elimination, one group was used for collection of sequential samples (1 ml) from the caudal vein into heparinized syringes 1, 3, 6, 12, and 24 h after tracer injection and then killed to determine tissue 109Cd levels. Additional blood and tissue samples were taken from animal groups killed 3 or 7 days after tracer injection alone (control) and 3 or 7 days after a single injection of radioinert CdCl2 (5 mg/0.2 ml water per kilogram body weight) followed by an injection of tracer 24 h before death (Cd pretreated). The dose of CdCl2 chosen is within the effective range for spermatotoxicity in rodents (1–8 mg/kg; [15, 16, 18–20]). All tissues were kept at 4°C during processing, and triplicate samples were taken for analysis. Whole blood and plasma were analyzed separately. Organ weights were determined for testes (plus epigonal organ), liver, kidney, spleen, heart, rectal gland, and brain. Additional samples were taken from skeletal muscle (body wall) and vas deferens. In some experiments, expressed contents of the vas (seminal fluid) and remaining tissue were analyzed separately. The spatial organization of spermatogenesis in the shark testis, procedures used for staging and dissection of tissues, and verification of the cellular composition of dissected samples by light and electron microscopy have been described previously [21–23]. In brief, after removal of the epigonal organ (EO; a lymphomyeloid tissue encapsulating the gonads in cartilaginous fishes), testes were cross-sectioned and dissected into the following zones: germinal (GZ), premeiotic (PrM), zone of degeneration (ZD), meiotic (M), and postmeiotic (PoM). For orientation, see diagrammatic cross section in Figure 1. To measure radioactivity, blood, plasma (100 µl each), and tissue (200–500 mg) were digested in 3 vol 2 N NaOH (w:v) at 55°C for ~20 h and analyzed using a Packard Autogamma 5000 Series gamma counter (Packard Instruments, Meriden, CT). Values were expressed as cpm/g wet weight, assuming a density of 1 g/ml for blood or plasma, and used to compute cpm/organ. Total blood volume was estimated at 6.8% and plasma at 5.5% body weight [24]. Total muscle volume was estimated as 75% of body weight. Total dose recovered was computed from the sum of measured and estimated radioactivity.

Tissue Cd Concentrations

To determine actual Cd levels, staged testicular tissues and liver, kidney, heart, and muscle were collected from untreated controls and from animals injected with CdCl2 (5 mg/kg body weight) 3 days before death as described above (3 per group); samples were stored at -70°C until analysis. Samples (200 mg) were weighed into acid-cleaned polytetrafluoroethylene vials and digested at 70°C in trace metal-grade concentrated nitric acid (Fisher, Pittsburgh, PA), which was then evaporated to dryness at 85°C. The residue was reconstituted in 3% nitric acid and filtered (0.45 µM). Cd was assayed on a Perkin-Elmer 5100 PC Atomic Absorption Spectrophotometer with Zeeman background correction equipped with a heated graphite atomizer furnace and programmer, an electrodeless discharge lamp, and an argon purge gas (Perkin-Elmer, Norwalk, CT). A matrix modifier (0.6 mg/ml Mg and 0.4 ml/ml Pd in 0.2% nitric acid) was added to the samples to stabilize Cd during the furnace char step. The linear range was 0–11 µg/L and the detection limit was 0.15 µg/L. For tissues measuring > 11 µg/L, samples were diluted 10-fold in 3% nitric acid. Certified atomic absorption Cd standards were from Perkin-Elmer. Values were expressed as µg Cd/g tissue.

Tissue Cd-Binding Activity

The Cd-hemoglobin affinity assay, a standard method of analysis based on 109Cd binding to heat-resistant tissue proteins, was used to estimate MT-like binding activity in staged testicular tissues [25]. Liver, kidney, heart, and muscle were used as controls. In brief, ~200 mg of frozen tissue was homogenized in 4 vol of homogenization buffer (HB; Tris-HCl, 10 mM, pH 7.4) and centrifuged at 10 000 x g for 10 min; the supernatant was boiled for 2 min. After centrifuging at 10 000 x g for 10 min, 100 µl of supernatant was incubated with the same volume of 109Cd solution (2 µg/ml CdCl2 and 1 µCi 109Cd in HB) for 10 min at room temperature. Addition of bovine hemoglobin (2% in 100 µl HB) followed by boiling for 2 min, cooling on ice for 5 min, and centrifugation (10 000 x g for 5 min) and then addition of hemoglobin, boiling, cooling and centrifuging again, separated free and bound fractions. Radioactivity in the supernatant (200 µl) was measured in a gamma counter (Searle Analytic, Des Plaines, IL). Blanks (no tissue extract) and total 109Cd (no tissue extract, no hemoglobin) were used as controls. To minimize isotope dilution by endogenous Cd (after Cd treatment) and to assure ligand availability, samples were diluted prior to assay such that bound 109Cd was no more than 20% of total added. Values were expressed as µg Cd bound/g tissue. MT II from rabbit liver (Sigma Chemical Company, St. Louis, MO) was used to standardize the assay.

Subcellular 109Cd Distribution

To determine the subcellular distribution of 109Cd in testes, like stages (GZ/PrM, M, PoM) were pooled from animals injected i.v. with a tracer dose of 109Cd 3 days (experiment 1) or 7 days (experiment 2) previously. Kidney was taken as control. Tissues were homogenized and cellular subfractions prepared using buffers and procedures essentially as described for characterizing steroid receptors in cytosolic and nuclear extracts of shark testis [26], with the exception that monothioglycerol was omitted from all buffers to avoid possible interference with Cd. Tissue aliquots (0.3–3.0 g) were homogenized in buffer H (1:3 w:v; 50 mM Tris HCl, 1 mM EDTA, 30% glycerol; pH 7.5) using a Polytron (Brinkman, Westbury, NY; three 15-sec bursts) and centrifuged at 1000 x g for l5 min to obtain a crude nuclear subfraction. The pellet was washed three times with buffer W (10 mM Tris HCl, 3 mM MgCl2, 0.25 M sucrose; pH 7.5), each time followed by centrifugation at 1000 x g for 15 min, and then extracted for 60 min with buffer H containing 0.7 M KCl (buffer E, original vol) and centrifuged at 100 000 x g for 60 min to obtain a salt-extractable nuclear fraction. The remaining pellet was designated the salt-resistant nuclear fraction. The 1000 x g supernatant was centrifuged at 100 000 x g for 60 min to obtain a cytosolic extract, and the remaining pellet was designated membranes (plus mitochondria). Radioactivity was measured in triplicate aliquots of each fraction. Values were expressed as cpm/g tissue.

Histology

To verify staging, tissue samples were fixed in Bouin's, embedded in polyester wax, sectioned at 6 µm, affixed to gelatin-coated slides, and then baked at 40°C overnight. Sections were stained for 5 min with a fluorescent nucleophilic dye, acridine orange (AO), to visualize distinctive patterns of cellular organization of spermatocysts at different stages. Images were viewed and recorded using an Olympus Fluoview Personal Confocal Microscope System (Olympus America Inc., Melville, NY) equipped with an Olympus IX70 inverted microscope and a UPLFL x10 objective (NA 0.30). The confocal system was controlled by the Fluoview 2.0.28 program for Windows NT. Illumination was provided by an argon/krypton laser at 488/568 nm and a 510-nm dichroic filter, a 550-nm long-pass emission filter, and a 6% neutral density filter.

Statistics

Statistical differences were examined by two-way ANOVA, one-way ANOVA, or Student's t-test (see figure legends; SigmaStat, Version 1.0, Jandel Scientific, San Rafael, CA). Where differences were detected, Student-Newman-Keuls multiple comparisons test was used to determine which values differed significantly (p < 0.05). In cases of non-normal distributions or unequal variances, log or square root transformations were performed.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Tissue Staging (Fig. 1)

Fluorescence confocal imaging of AO-stained sections verified staging of dissected tissues. As shown in Figure 1, A–E, stage of development of spermatocysts was identifiable by cyst diameter; the presence or absence of a lumen; and the approximate number, ratio, and relative position of germ cell vs. Sertoli cell nuclei. All spermatocysts at a given distance from GZ were in the same stage of development. Dissected regions designated PrM were most heterogeneous, as there are 13 spermatogonial generations in Squalus [4, 7].

Elimination of 109Cd from the Peripheral Circulation (Fig. 2; Table 1)

Within 1 h of an i.v. injection of a single tracer dose of 109Cd, radioactivity recovered in the peripheral blood was 31% of the injected dose, and levels decreased progressively thereafter to 4% of the injected dose by Day 7 (Table 1). Elimination curves for blood and plasma were biphasic and, when plotted as log concentration vs. time, indicated two components: a fast component with a half-time of 3.2 h in blood and 1.6 h in plasma, and a slow component with half-times of 150 h and 50 h in blood and plasma, respectively (Fig. 2). Initially (1 h postinjection), the blood : plasma ratio was < 1.0, indicating that the majority of tracer was in the fluid compartment; however, 6 h and 7 days after injection, the blood:plasma ratio had increased to 3:1 and 24:1, respectively, indicating tracer retention in the cellular compartment.



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  		FIG. 2. Blood and plasma elimination of 109Cd. Blood was sampled 1, 3, 6, 12, and 24 h after 109Cd injection, and animals were killed at the 24-h time point for tissue analysis (n = 3). Additional samples were taken from animals killed 3 and 7 days after tracer injection (n = 3 each). Samples were assayed in triplicate. Values are the mean and SEM of 3 animals. Biphasic curves were transformed to linear plots and t1/2 was calculated as the time taken to reach 50% of the time zero value.


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  		TABLE 1. Organ weights and body distribution (% bw = % body weight) of 109Cd in male dogfish shark (Squalus acanthias) 1, 3, and 7 days after tracer administration, and effects of Cd pretreatment.

109Cd Distribution in the Testis and Control Tissues (Fig. 3; Table 1)

By 24 h postinjection, < 1% of the injected dose had accumulated in testis, an amount lower than testicular mass as a percentage of body weight (< 2%); however, the same fraction of administered 109Cd was retained in the testis for at least 7 days postinjection (Table 1). Significantly, intratesticular 109Cd concentrations were stage dependent, with highest levels in regions with stem cells (GZ) and spermatogonial-stage cysts (PrM) and progressively lower levels in regions with more mature developmental stages. This resulted in a decreasing intratesticular 109Cd gradient (immature > mature). Absolute levels of radiolabeled Cd increased in all testicular regions between 1 and 3 days and between 1 and 7 days postinjection, but the increase was greatest in GZ and PrM regions. Thus, when measured as the fold difference between least mature (GZ) and most mature (PoM) regions at 1, 3, and 7 days postinjection, the 109Cd gradient was 3-, 4.7-, and 3.9-fold at the respective time points. Regardless of position in the gradient, testicular 109Cd levels 3 and 7 days after tracer injection always exceeded those in plasma. The 109Cd-concentrating potential of GZ, when expressed as tissue:plasma ratios, was 2:1, 7:1, and 71:1, respectively, at 1, 3, and 7 days postinjection (compare Figs. 2 and 3A).



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  		FIG. 3. Stage-related distribution of 109Cd in A) male reproductive organs and B) control tissues. Note scale difference between A and B. 109Cd levels were determined 1, 3, or 7 days after tracer injection. VAS, vas deferens; KID, kidney; LIV, liver; HRT, heart; MUS, muscle. Values represent the mean and SEM of 3 animals, each determined in triplicate. Intratesticular values were analyzed by two-way ANOVA as a function of region (p < 0.001) and time (p < 0.001), and significant differences were determined by the Student-Newman-Keuls post hoc comparison test (p < 0.05). For a given day posttracer, regions with different letters differ significantly: Day 1, a–c; Day 3, k–m; Day 7, x–z. For a given region, significant differences between days are indicated by an asterisk (*). Values for EO, VAS, and nonreproductive control tissues were analyzed separately as a function of time after tracer (one-way ANOVA).

109Cd levels in the EO were higher than in adjacent PoM tissues. Although 109Cd levels in the vas deferens were similar to those in testicular regions with elongated spermatids (PoM), < 3% of the total radioactivity was recovered from the expressed seminal fluid (containing sperm bundles at this time of year). Compared to levels in the testis, accumulated levels of 109Cd/g were higher in kidney, liver, spleen, heart, and rectal gland and lower in skeletal muscle and brain (Fig. 3B; Table 1). The liver accounted for the highest percentage of injected dose, followed by skeletal muscle, blood, and kidney (Table 1). Only 68% of injected dose was accounted for after 24 h, indicating excretion or the presence of tracer in tissues not measured (e.g, urine, bile, skin). Although the mean body weight of animals killed at 1 day postinjection was somewhat higher than those at Days 3 and 7 (p < 0.05), there were no significant time-related differences when extragonadal tissue concentrations, fraction of injected dose, or percentage dose recovered were compared at the three time points.

Effects of Pretreatment with Radioinert CdCl2 on 109Cd Distribution in Testis and Control Tissues (Fig. 4; Table 1)

Pretreatment with radioinert CdCl2 2 or 6 days before tracer injection significantly reduced 24-h accumulations of 109Cd in all testicular zones except GZ but did not alter the stage-related pattern (Fig. 4A). Prior Cd exposure did not affect the total percentage of tracer accounted for or the fractional recovery and the measured 109Cd concentrations in most extragonadal tissues (Fig. 4B; Table 1). Exceptions were liver and plasma. CdCl2 pretreatment significantly reduced hepatic tracer accumulation (Fig. 4B) and elevated 109Cd levels in plasma from 1960 ± 127 cpm/g in controls to 5368 ± 1948 cpm/g at 3 days and 4743 ± 569 cpm/g at 7 days after CdCl2, but it did not change whole blood concentrations (6176 ± 1083, 5150 ± 1345, and 4198 ± 485 cpm/g in, respectively, controls and at 3 and 7 days after CdCl2). This implies a shift of Cd-binding components from blood cells to plasma.



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  		FIG. 4. Effects of Cd pretreatment on stage-related distribution of 109Cd in A) male reproductive organs and B) control tissues. Note scale difference between A and B. Animals received a single injection of CdCl2 3 or 7 days before being killed and a tracer dose of 109Cd during the last 24 h. Values represent the mean and SEM of 3 animals, each determined in triplicate. Statistical analysis was performed as described for Figure 3.

Radioinert Cd Levels in Testis and Control Tissues (Fig. 5)

When actual Cd levels were measured by atomic absorption spectrophotometry in tissues of wild-caught animals, the intratesticular gradient was like that seen after 109Cd injection (immature > mature), but zonal differences measured by ANOVA were not significant due to high animal-to-animal variability and measurements close to detection limits. A single injection of CdCl2 3 days prior to death dramatically increased testicular Cd. Although the fold increase varied by stage (GZ, 166-fold; PrM, 216-fold; M, 545-fold; PoM, 171-fold), the stage-related pattern (immature > mature) and the magnitude of the gradient (3- to 4-fold) remained the same (compare Figs. 3 and 5). In comparison to levels in testis, basal Cd levels in nongonadal tissues were much higher, but the increase after CdCl2 treatment was of lower magnitude (29-, 72-, and 105-fold in liver, heart, and kidney, respectively).



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  		FIG. 5. Differences in Cd concentrations and effects of Cd pretreatment in A) the testis by stage and B) control tissues. Animals were untreated or were given a single injection of CdCl2 3 days before being killed. Values represent the mean and SEM of 3 animals, each determined in triplicate. A) Significance was determined by two-way ANOVA as a function of testicular region (p < 0.001), treatment (p < 0.001), and interaction between the two variables (p < 0.001). Regional differences in a given treatment group were determined by a post hoc comparison test (p < 0.05), and those with different letters differed significantly: untreated, a; Cd treated, k–m. Within a given region, differences between untreated and Cd-treated animals are indicated by an asterisk (*). B) Statistical comparisons were not made between tissues. For a given tissue, differences between control and treated groups were determined by Student's t-test (*, p < 0.05).

Tissue 109Cd-Binding Activity (Fig. 6)

A standard MT assay was used to measure basal and Cd-induced changes in heat-stable Cd-binding activity. Binding activity was stage dependent (PrM = PoM > GZ = M) in untreated animals, but zonal differences did not correspond to those predicted by tracer or atomic absorption spectrophotometry analysis. Three days after a single injection of radioinert CdCl2, MT-like activity increased in all but M stages (Fig. 6), and the response in GZ (4-fold) was of greater magnitude than in PrM and PoM stages (2.5- and 1.8-fold; Fig. 6). In untreated animals, MT-like binding activity in kidney and liver was 3 to 15 times higher than in testis, but CdCl2 induced similar fold increases in all three tissues (3- to 4-fold). Interestingly, induction of the MT-like component was greatest in the heart (23-fold). Comparison of Figures 5 and 6 shows that the MT-like protein accounted for only a small fraction of total bound Cd, regardless of tissue type or treatment.



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  		FIG. 6. Differences in 109Cd-binding activity and effect of Cd pretreatment in A) staged testicular tissues and B) control tissues. Animals were untreated (n = 8) or were given a single injection of CdCl2 3 days before being killed (n = 3). Dissected tissues were processed to obtain extracts, and 109Cd-binding activity was analyzed in the heat-resistant fraction (see Materials and Methods). Values were expressed in µg 109Cd bound/g original tissue wet weight and represent the mean of all animals per treatment group with duplicate determinations per animal. A) Data were analyzed by two-way ANOVA to determine significance by stage (p < 0.001) and treatment (p < 0.001). Regional differences in a given treatment group were determined by a post hoc comparison test (p < 0.05), and those with different letters differed significantly: untreated, a, b; Cd treated, k, l. For a given region, differences between control and treated groups are indicated by an asterisk (*). In B, different tissue types were not compared statistically. For a given tissue, differences between control and treated groups were determined by Student's t-test (*, p < 0.05).

Subcellular Distribution of 109Cd (Fig. 7)

After injection of tracer, radioactivity (cpm/g) in cytosolic, nuclear, and membrane subfractions reflected levels in whole homogenates and whole tissues (compare Figs. 3 and 7). Regardless of tissue type or stage, proportionally more 109Cd was recovered from cytosol (~75%) than from other subfractions; however, fractional recovery of radioactivity from cell nuclei in GZ/PrM stages (14% and 13%, experiments 1 and 2, respectively) was > 2-fold greater than radioactivity in this subfraction in other stages or in kidney (< 6%, both experiments).



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  		FIG. 7. Stage-related differences in the subcellular distribution of 109Cd. Animals received a single injection of 109Cd and were killed after 3 days (n = 1, experiment 1) or 7 days (n = 2, experiment 2). Dissected tissues were processed to salt-resistant and salt-extractable nuclear fractions, cytosol, and membranes (plus mitochondria). Radioactivity was measured in triplicate in each fraction. Values for each fraction were expressed as cpm/g tissue.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Data reported here reveal an intratesticular stage-related gradient of Cd uptake and retention. Cd accumulated preferentially in GZ, with decreasing levels in succeeding developmental stages. The direction (immature > mature) and magnitude (3- to 5-fold) of the intratesticular gradient were essentially the same whether Cd was measured as radioactivity after a single injection of 109Cd or by atomic absorption spectrophotometry after administration of radioinert CdCl2. Moreover, the same pattern was seen in untreated controls, indicating that the gradient was not a treatment artifact and implying that animals are exposed to Cd in their environment. Cd occurs in nature in ores and minerals, but levels in seawater are generally low [12, 13]. Because Cd undergoes bioaccumulation and biomagnification in the food chain, and sharks are apex feeders, tissue Cd is likely to be dietary in origin. Identifying the source of this exposure would be difficult, however, because Squalus migrates annually between the North and South Atlantic.

The observed intratesticular Cd gradient cannot be ascribed to a dilution effect related to the pattern of vascular flow. In sharks, blood from the genital artery first perfuses the EO and then passes successively from mature to immature regions [27], which is the reverse of the gradient. Also, in contrast to the rapid elimination of 109Cd from plasma, testicular 109Cd increases progressively in the 3-day interval after injection and then persists unchanged for an additional 4 days. Long retention times are seen in all tissues, despite tissue-specific differences in kinetics and final Cd concentrations, indicating a common high-affinity binding mechanism. When the power of this mechanism in different shark tissues is estimated as tissue:plasma ratios 7 days after tracer injection, binding in GZ and that in liver are equivalent (71:1 and 87:1, respectively) but not as powerful as in kidney (381:1). Even at the shallow end of the gradient, PoM-stage tissues are able to concentrate 109Cd to levels 18-fold higher than in plasma. Consistent with our results, testicular Cd accumulation is 5-fold lower in adult rats than in 4-day-old animals [28]. Paradoxically, 109Cd accumulation in the adult rat testis is maximal in elongated spermatids, with spermatogonia showing only a very low incorporation [15].

MT are a family of low molecular weight, cysteine-rich proteins that bind metal ions with high affinity [17]. Several studies have reported a relationship between MT concentrations and differences in tissue, strain, and species in Cd-accumulation and -storage potentials [28–32]. MT has been partially characterized in the liver of the spotted dogfish shark (Scyliorhinus), and trace amounts were detected in the testis [33–36]. Although the binding component measured in our assays was not definitively identified as MT, it is heat stable, comigrates with rabbit liver MT II on SDS-PAGE gels (< 10 kDa), and gives the same anomalous staining with Coomassie blue (indicating a low content of aromatic amino acids; unpublished results). This protein also resembles MT in its Cd inducibility. Indeed, a single CdCl2 injection up-regulates MT-like activity to the same extent in testis as in kidney and liver. This is surprising because MT protein and mRNA in rodent testis are refractory to Cd induction [29, 31, 37]. Although the distribution of MT-like activity in Squalus testis is stage related (PrM = PoM > GZ = M), the pattern differs from that predicted by tissue Cd levels. Moreover, increased levels of MT-like activity after CdCl2 pretreatment do not affect the 109Cd-accumulating gradient, nor is the fold increase (4-fold) sufficient to account for the increase in testicular Cd (> 160-fold). Although we cannot rule out the possibility that the standard Cd hemoglobin assay underestimates the number of Cd-binding sites (due to occupancy by radioinert Cd), the data are consistent with the view that MT is not the primary determinant of stage-related Cd accumulation.

To investigate the presence of Cd-binding components other than MT, which is mainly cytosolic, 109Cd was analyzed in different tissue subfractions after tracer administration in vivo. Although cytosolic binding components are quantitatively most important and can account for the stage-related accumulation pattern, a second nuclear-associated component is detected in greater amounts in immature than in mature stages. In rodents, the amount of Cd associated with the nucleus increases with increasing extracellular Cd, and this has been interpreted as induction of metal-binding nuclear proteins and/or translocation of metal-binding cytosolic proteins [38]. Alternatively, nuclear-associated Cd may displace zinc in polymerases, other metalloenzymes, or transcription factors, all of which are present in greater amounts in nuclei of proliferating cells [39]. It may be relevant here that androgen and estrogen receptors, which are zinc finger proteins, are concentrated in PrM regions of shark testis [23, 26]. Finally, because Cd has a high affinity for nucleic acids and accumulates within the nuclear compartment even when present in trace amounts [39], high levels of nuclear-associated Cd in the GZ/PrM region may reflect the higher percentage of cells with replicated DNA.

While it is tempting to interpret the intratesticular Cd gradient in shark testis as due to developmental programming of genes encoding the number of Cd-binding molecules per cell, there are alternative explanations. One possibility is that it reflects stage-related differences in the number, ratio, and size of germ cells vs. Sertoli cells in each region ([9]; see Fig. 1). Another explanation is that Cd accumulation decreases with development of the blood-testis barrier. In shark testis, tight junctions form between adjacent Sertoli cells when spermatocysts are in late spermatogonial or early spermatocyte stages of development [40], and exclusion of dextran-rhodamine (10 000 Mr) from the germinal compartment is observed in isolated M and PoM, but not PrM, spermatocysts [10]. Higher levels of Cd in immature compared to mature rat testes, and increased Cd accumulation after Cd pretreatment, have been attributed to the status of the blood-testis barrier [28, 41, 42]. Nonetheless, the barrier per se cannot account for the Cd gradient in shark testis. Although Cd exposure in vivo and in vitro perturbs the blood-testis barrier in sharks, as measured by dextran-rhodamine exclusion in PoM cysts [10], it does not alter the pattern of Cd accumulation.

It is generally agreed that Cd uptake and accumulation are the first steps in an effector pathway leading to a biological or toxic response, but the relationship between the amount of Cd bound and tissue sensitivity or responsiveness to Cd is still debated. The problem is complicated by Cd induction of factors like MT. MT has been variously reported to reduce [43–45], enhance [11, 46], or have no role [32, 47, 48] in susceptibility to metal toxicity. Initial results in sharks indicate that a single injection of CdCl2 increases the percentage of cysts with apoptotic germ cells in PrM stages where accumulation is maximal [10, 49]. Also, data in this report show that MT-like binding activity increases to a greater extent in GZ and PrM than in later stages. The complete lack of responsiveness to MT induction by Cd in M stages may be due to a general shutdown of transcription that is known to occur in spermatocytes in rodents [50]. High levels of type I and II MT mRNA have been localized in germ cells in mouse testis [31] but, in contrast to the stage-related pattern of MT-like protein in sharks, mRNA levels are highest in spermatocytes and round spermatids [31, 51].

Although further work is required to characterize the intratesticular Cd-binding mechanism and to determine its localization in germ cells or Sertoli cells, we conclude that the shark testis model has utility for stage-by-stage analysis of spermatotoxicant accumulation and effect.


   ACKNOWLEDGMENTS
 
We thank David Miller for invaluable advice and Leon McClusky for histological preparation of tissues. Work was carried out in part at the Mount Desert Island Biological Laboratory (Salsbury Cove, ME).


   FOOTNOTES
 
1 This research was supported by a grant from the National Institutes of Environmental Health (NIEHS P42 ES-07381). Back

2 Correspondence: G.V. Callard, Department of Biology, Boston University, 5 Cummington Street, Boston, MA 02215. FAX: 617 353 6340; gvc{at}bio.bu.edu Back

Accepted: August 18, 1998.

Received: April 8, 1998.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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