Animals

Crl:CD1(ICR) (CD1) outbred mice were obtained from Charles River Laboratories (Wilmington, MA) or an in-house vivarium. WBB6F1/J-Kit^W/Kit^W-V (W/W^v) mice were obtained from an in house vivarium or purchased from Jackson Laboratories (Bar Harbor, ME). Animal experimentation and procedures were approved by the Washington State University Institutional Animal Care and Use Committee. Animals were treated humanely in accordance with the standards of care outlined by the NIH Guide for the Care and Use of Laboratory Animals.

DEHP Treatment and Breeding to Obtain F1–F4 Offspring

Postnatal Day (P) 60–64 (P60–64) CD1 females were continuously paired with age-matched CD1 males for generating offspring. Females were examined twice daily for the presence of vaginal plugs, the presence of which was set as E0. Twenty vaginal plug-positive dams were gavaged from E7 to E14 of gestation with DEHP (500 mg/kg body weight/day) (Sigma-Aldrich Chemical Co., St. Louis, MO) in 200 μl of superrefined corn oil (Croda Inc., Edison, NJ) (DEHP-treated group) or gavaged with 200 μl of corn oil alone (vehicle control group). A review [38] citing the comparison of the measurement of phthalate metabolites in amniotic fluids in rodents [39] and humans [3] indicated that the dose range including a 300 mg/kg/day dose in animal studies is relevant to understanding TDS in humans and the animal dosages are not excessively high as originally thought. The treated gestating dams were considered the F0 generation (F0) dams, and the pups born from the F0 dams were labeled as the F1 generation (F1) offspring. The F1 offspring were used in three breeding paradigms to follow inheritance through the maternal, paternal, and double-cross lines. Paternal lines were generated by breeding males from the treated groups (both DEHP-treated and vehicle control) with age-matched nontreated CD1 females. Maternal lines were generated by breeding females from the treated groups with age-matched nontreated CD1 males. For the double-cross lines, males were bred to females, both from the same treated group. Each paradigm was repeated to produce F2 to F4 offspring. No inbreeding was performed.

Embryonic Survival and Weaning Rates

The number of implantation sites on uterine horns, representing the number of embryos embedded in the uterine wall, was determined from the F0 dams at the day of pup weaning using a stereomicroscope (Leica MZ 9.5; Leica, Wetzlar, Germany). The litter size was noted on P1, the body weights were measured at P10, and the number of pups successfully reared to weaning was noted on the day of weaning (P21).

Neonatal and Pubertal Period Analysis

Anogenital distance (AGD) was measured on neonatal pups at P1 by two independent researchers. The distance from the urethral opening to the ventral opening of the anus was measured in millimeters under a stereomicroscope (Leica MZ 9.5). Pups at P10 were anesthetized and nipple positions were determined by measuring the distance of each nipple to the umbilical stump. The nipples were numbered in a pattern beginning with the right foremost cranial nipple labeled nipple 1, the right axillary nipple labeled nipple 2, and continuing down to the rightmost caudal nipple labeled nipple 5. The order was reversed on the left side with the leftmost caudal nipple labeled nipple 6, the inguinal nipple labeled nipple 8, and continuing up to the left foremost cranial nipple, labeled nipple 10. Pubertal onset occurs around the time when the penis detaches from the prepuce in male pups and was determined by retracting the prepuce manually using gentle pressure beginning at P20 onward.

Testicular Tissue Collection and Determination of Testicular Abnormalities

Male mice were killed at P30 and P120 for F1–F4 generations and P360 for F4 generation and their testes were collected, fixed in Bouin solution for 1–5 h or 4% paraformaldehyde (PFA) with 0.25% glutaraldehyde for 4–6 h at 4°C, and embedded in paraffin. Four-micrometer-thick serial sections were cut and mounted on poly-l-lysine slides. To examine testicular morphology, sections were deparaffinized, stained with hematoxylin and eosin, and examined using a Leitz DMRB microscope (Leica). Digital images were obtained by using a Magnafire digital camera (Optronics, Goleta, CA). To determine the percentage of abnormal tubules present in the testes, tubules having moderate to severe germ cell disorganization and the total number of tubules were counted in testicular sections. To obtain good sampling across the entire testis, a minimum of four sections spaced 50 μm apart were counted. This was repeated for at least three animals.

Detection of Apoptosis by TUNEL Assay

Testicular sections fixed in 4% PFA with 0.25% glutaraldehyde were used for the detection of apoptotic cells by TUNEL assay (Promega Corporation, Madison, WI). This assay identifies fragmented DNA by the terminal deoxynucleotidyl transferase (TdT)-mediated incorporation of fluorescein-12-dUTP at the 3′ hydroxyl end of DNA. For a positive control, tissue sections were pretreated with deoxyribonuclease I, and for a negative control, tissue sections were not incubated with TdT. Apoptotic cells were visualized by green fluorescence using a microscope. Tubules with four or more apoptotic cells (representing apoptotic tubules) and the total number of tubules were counted to quantify the results.

Sperm Count and Determination of Sperm Motility

Sperm was collected from the caudal epididymis of F1–F4 male offspring at P120. The excised epididymides were cut in 2-mm intervals (five cuts) and placed in F12 media (1 ml) containing 0.1% bovine serum albumin prewarmed to 37°C and incubated for 15 min to facilitate sperm transmigration from the epididymis. Fresh sperm were loaded into a hemacytometer and counted. Each chamber was counted twice and averaged. A prewarmed glass slide was loaded with fresh sperm, coverslipped, and examined by phase contrast light microscopy to determine sperm motility. Sperm motility was scored in five randomly chosen fields for two samples from each animal. The number of sperm exhibiting forward and progressive activity and the total number of sperm were counted to obtain the percentage sperm motility. Sperm clumps, detached heads, and free tails were not counted.

Spermatogonial Transplantation

Three F3 male offspring at P24–P31 from the DEHP-treated group in the double-cross line provided the donor germ cells for spermatogonial transplantation. Germ cells from two age-matched untreated CD1 male mice served as controls. As described [40], the testes were detunicated, tubules were digested with enzymes, and single-cell suspensions were obtained. The donor germ cells were counted using a hemacytometer and resuspended at a concentration of 10⁷ cells/ml in minimal essential media α with 0.03% trypan blue dye (Invitrogen Corporation, Carlsbad, CA). Donor germ cells were transplanted into testes of at least three W/W^V mice per donor animal. A small hole was made in the connective tissue enclosing the efferent ducts of the recipient mouse and a glass needle containing 7 μl (70 000 cells) of the donor germ cell suspension was inserted into the rete testis through the efferent ducts for the injection. After injection, the animals were allowed to recover for 70 days. By this time, all stages of germ cells, including meiotic and postmeiotic germ cells, were represented in the recipient tubules injected with control SSCs.

Posttransplantation Recovery Analysis

The recipient WW^V testicular tissues and noninjected WW^V testicular tissues were fixed in Bouin solution for 2 h and embedded in paraffin, and 4-μm-thick serial sections were cut. To determine the donor-derived germ cell recovery, the tubules containing round or elongated spermatids, more than four sections separated by 50 μm per testis from at least three animals, were stained with hematoxylin and eosin and examined using a Leitz DMRB microscope, and digital images were obtained using a Magnafire digital camera. To further determine the germ cell types present in the recovered tubules, immunohistochemistry or immunofluorescence was performed using antibodies for DDX4, γH2AX, STRA8, and DMRT1.

Immunohistochemistry and Immunofluorescence

For immunohistochemistry, after a sodium citrate antigen retrieval step, tissue sections were incubated overnight with rabbit anti-STRA8 peptide antibody (1:2500 dilution) or rabbit anti-DDX4 peptide antibody (1:300 dilution; Abcam, Cambridge, MA), washed in PBS, and incubated with biotinylated anti-rabbit IgG (1:300 dilution; Vector Laboratories, Burlingame, CA), followed by streptavidin-peroxidase reaction and incubation with 3,3′-diaminobenzidine tetrahydrochloride (Invitrogen Corporation). For immunofluorescence, the tissue sections were further permeabilized with 0.2% Triton X-100 in PBS for 45 min. The rabbit polyclonal anti-DMRT1 antibody (1:300 dilution) incubation was followed by anti-rabbit IgG conjugated to Alexafluor-555 (1:1000 dilution; Invitrogen Corporation) secondary antibody. The mouse monoclonal anti-γH2AX peptide antibody (1:750 dilution; Millipore Corporation, Billerica, MA) incubation was followed by the use of Mouse on Mouse Basic Kit (Vector). Coverslips were mounted with ProLong Gold antifade reagent with 4′,6-diamidino-2-phenylindole (Invitrogen Corporation) on sections and examined by fluorescent microscopy (Leitz DMRB microscope).

Statistical Significance

Statistical significance was determined by a pooled t-test or one-way ANOVA, followed by pairwise comparison of the means using the Student t method (JMP 9; SAS Institute Inc., Cary, NC). The P value of <0.05 was considered significant. We made a note of P values between 0.05 and <0.1 in the text.

DEHP Effect on Embryo Survival and Weaning Rates in CD1 Outbred Mice

We established the effect of a 500 mg/kg/day dose of DEHP between E7 and E14 on the embryo survival of the CD1 outbred strain. This window of treatment for CD1 mice had not been conducted previously. The number of implantation sites and litter sizes were determined for the F0, F1, and F2 dams. We found that the E7–E14 exposure of DEHP did not significantly affect the number of implantation sites in the DEHP-treated F0 dams (Fig. 1A). This was expected, as the treatment started 7 days after implantation. However, the litter size of the DEHP-treated F0 dams compared to vehicle control dams was different at P = 0.0667, showing a trend of 17.2% decrease in the litter size for DEHP-treated F0 dams (Fig. 1B). After birth, there was no significant difference at P < 0.05 in the wean rate of F1 pups between the DEHP-treated dams (n = 6) and vehicle-treated dams (n = 3). The neonatal survival of F1 offspring was 100%. For F2 and F3 embryo and neonatal survival, there were no significant differences in the number of implantation sites, the litter size of dams, and the wean rate of pups between the DEHP-treated and vehicle-treated groups for the paternal, maternal, and double-cross lines at P < 0.05 (Table 1). Thus, the embryo survival rate returned to normal for F2 and F3 embryos, indicating that the embryo survival rate is not transgenerational.

FIG. 1

Implantation sites and litter size for F0 dams, and AGDs and body weights of F1 offspring. Time-pregnant dams were treated with 500 mg/kg body weight/day DEHP or corn oil vehicle (CO) from E7 to E14. Individual value plots are shown for the number of implantation sites in F0 dams (A), the number of live pups born per litter for F0 dams (B), AGD measurements in millimeters for F1 pups at P1 (C), and the body weights in grams for F1 pups at P10 (D) from DEHP-treated or corn oil vehicle (CO)-treated dams. The number (n) in the parentheses indicates the number of dams for A and B and the number of male offspring for C and D. Values are mean ± SEM. Pooled t-tests were conducted for the control and DEHP values; ***P < 0.001, no asterisk indicates P > 0.05.

TABLE 1

Implantation sites, litter size, and weaned rate for F1 and F2 dams.^a

a 

Pooled t-tests were conducted, and P values for the DEHP group to the control group is not significant at P < 0.05.

b 

M, maternal line; P, paternal line; DC, double-cross line.

c 

CO, corn oil vehicle control.

DEHP Effect on Anogenital Distance, Body Weight, Nipple Retention, and Pubertal Onset

Fetal and lactational DEHP exposure that includes late embryonic stage E16–E20 has been found to decrease male AGD and increase the incidence of males retaining nipples/areolas in rats [12–14]. To examine if mouse embryos exposed to DEHP from E7 to E14 have similar alterations, the AGD at P1 and nipple/areola retention at P10 were examined. The F1 male offspring at P1 from the DEHP-treated group exhibited a significant decrease (P < 0.0001) in AGD when compared to their vehicle control male counterpart (Fig. 1C) or nontreated control CD1 pups (data not shown). The average AGD for the vehicle control group was 1.90 mm and for the DEHP group was 1.67 mm at P1. Moreover, the body weights of F1 male offspring at P10 from the DEHP group were greater (average of 7.81 g) than those of the vehicle-treated group (average of 5.63 g) (Fig. 1D). Additionally, we detected alopecia/xerosis in 83.4% of F1 offspring in the DEHP group (n = 31), whereas no incident was observed for the control group (n = 17). However, the decrease in male AGDs, the increase in body weights, and the occurrence of alopecia/xerosis were not passed to F2 and F3 offspring from the maternal, paternal, and double-cross lines, evaluated on 21–43 male offspring for each breeding line/treatment group.

Supernumerary nipples were observed with a 16.1% frequency in F1 male pups at P10 from the DEHP-treated group (n = 31) compared to no supernumerary nipples in the vehicle-treated (n = 17) male pups. The retained nipples corresponded to nipple positions 3, 4, and 8 (as described in Materials and Methods). Examining the positions individually, 9.7% of pups had supernumerary nipples at both positions 3 and 4, and 12.9% of pups had supernumerary nipples at position 8. However, the incidence of retained nipples was not transmitted to F2 and F3 offspring males (n = 21–43) for any breeding line or treatment group.

To determine if DEHP exposure disrupted pubertal onset, the penis detachment from the prepuce was examined in males starting at P20. The average day of pubertal onset was P24.7 in the controls. The F1 offspring from DEHP-exposed dams had significantly delayed (P < 0.05) pubertal onset compared to controls (Fig. 2A). For F2 and F3 offspring, no significant difference at P < 0.05 was detected in pubertal onset between offspring of the DEHP-exposed and vehicle-exposed dams in the maternal line (Fig. 2, B and C). In contrast, the F2 and F3 offspring from the DEHP-exposed group had a significant delay (P < 0.05 to < 0.0001) in pubertal onset in the paternal and the double-cross lines compared to their vehicle counterparts (Fig. 2, B and C). These results show that DEHP decreased AGD, increased nipple retention, increased body weights, and caused alopecia/xerosis in F1 males, but these changes remained only in F1 offspring and did not transmit to F2 and F3 offspring. In contrast, male pubertal onset was delayed in F1 offspring and in F2 and F3 offspring of the paternal and double-cross lines, although the delay of pubertal onset was on average less than a day, relatively small for an outbred strain.

FIG. 2

Male pubertal onset of F1, F2, and F3 male offspring. The age of male pups was noted when prepuce detachment occurred. The average age of pubertal onset for F1, F2, and F3 male offspring in the vehicle control group was P24.7. Changes in days of the pubertal onset for F1 (A), F2 (B), and F3 (C) male pups from the corn oil vehicle (CO) and DEHP groups for the maternal (M), paternal (P), and double-cross (DC) lines were graphed. The number (n) in the parentheses indicates the number of male offspring for the CO and DEHP groups. Values are mean ± SEM. Pooled t-tests were conducted for the control and DEHP values; ***P < 0.001, *P < 0.05, no asterisk indicates P > 0.05.

DEHP Increased Abnormal Seminiferous Tubule Morphology in F1–F4 Offspring

Hematoxylin and eosin staining of testicular cross sections of F1–F3 offspring at P120 in the DEHP-treated group for the maternal, paternal, and double-cross lines revealed morphological abnormalities. The weights of testes from the control and DEHP group animals at P120 in the F2 and F3 generations were not significantly different at P < 0.05 (data not shown). For the vehicle control group, the testes of F1 (Fig. 3, A, C, and E), F2 (Fig. 4, A and E), F3 (Fig. 4, I and M), and F4 (Fig. 4, Q and R) offspring at P120 showed normal germ cell associations in the seminiferous tubules, with the different stages of the seminiferous epithelium cycle represented. In contrast, testes of F1 (Fig. 3, B, D, and F), F2 (Fig. 4, B–D and F–H), and F3 (Fig. 4, J–L and N–P) offspring at P120 from the DEHP-treated group in all three breeding lines had moderate to severe testicular abnormalities. On average, 27.1% (the range in the F2 generation was 18.8–34.2%, and in the F3 generation was 22.3–41.0%) of their seminiferous tubule sections had abnormalities in the F2 and F3 generations such that they could not be assigned to specific stages of the seminiferous epithelium. Tubules with moderate and severe abnormalities were easily identifiable, with one or more characteristics such as massive cell sloughing into the center of tubules (Fig. 3D and Fig. 4, D, N, and P), no lumen (Fig. 3, D and F, and Fig. 4, D, F, G, H, K, N, and P), a complete loss of germ cell organization (Fig. 3F and Fig. 4, F, G, H, K, and N), and large vacuoles from apparent bulk germ cell loss (arrow, Fig. 4, C, J and N). Also, a small number of tubules had areas where layers of germ cells were missing (oblong circle, Fig. 4, G, J, L, and O). The germ cells missing were round spermatids (Fig. 4, j‘ and l‘), leptotene (Fig. 4o‘) and pachytene spermatocytes (Fig. 4g‘). Likewise, testes of F4 offspring at P120 from the DEHP-treated group in the double-cross line (Fig. 4, S and T) had testicular morphologies similar to those from F1–F3 offspring from all three breeding lines.

FIG. 3

Testicular cross sections from F1 offspring. Hematoxylin and eosin-stained testicular cross sections for F1 offspring at P120 after embryonic exposure to corn oil vehicle (CO; A, C, and E) and DEHP (B, D, and F). The stages of seminiferous epithelial cycle are noted in control sections (A, C, and E). Arrowheads indicate small vacuoles (B). Bar = 50 μm. Photomicrographs are representative of n ≥ 6 offspring for both the vehicle control and DEHP groups from two sets of embryonic exposures.

FIG. 4

Testicular cross sections from F2, F3, and F4 offspring at P120. Hematoxylin and eosin-stained testicular cross sections for the F2 and F3 offspring at P120 in the control vehicle group (CO; A, E, I, and M) and DEHP group in the maternal (M; B, F, J, and N), paternal (P; C, G, K, and O), and double-cross (DC; D, H, L, and P) lines and for the F4 offspring at P120 in the control vehicle group (Q and R) and DEHP group (S and T) in the double-cross line (DC) are shown. The stages of seminiferous epithelial cycle are noted in control sections (A, E, I, M, Q, and R). Arrowheads indicate areas where small vacuoles were observed. Arrows indicate areas where large vacuoles were observed. Oblong circles are magnified in g‘, j‘, l‘, and o‘, showing areas with germ cells missing. Bar = 50 μm. Photomicrographs are representative of n ≥ 6 offspring for the F2 and F3 generations and three breeding lines from two sets of embryonic exposures and n ≥ 3 offspring for the F4 generation.

To determine if the testicular abnormalities observed at P120 were progressively worse with aging, we examined the testes from F4 offspring at P360 in the double-cross line. Similar to testes from the P120 mice, seminiferous tubules of animals in the corn oil group had distinguishable stages of the seminiferous epithelium cycle (Fig. 5, A, D, and G). In contrast, testes from mice at P360 in the DEHP group showed abnormal testicular morphologies, much more severe than those observed at P120. Abnormal tubules showed large vacuoles (arrows, Fig. 5, E, F, and I), loss of germ cell organization (Fig. 5, B and H), and presence of multinucleated cells (Fig. 5, B and F, box and insets b‘ and f‘), which are thought to be precursors to cells that may become cancerous. Some severely abnormal tubules contained only a few layers of germ cells (Fig. 5I) and some had masses of unknown cell types (Fig. 5C). Epididymides of F4 offspring from the DEHP group showed round germ cells and multinucleated germ cells (Fig. 5, K and L), which should not be present in epididymides in normal condition.

FIG. 5

Testicular and epididymal cross sections from F4 offspring at P360. Hematoxylin and eosin-stained testicular and epididymal cross sections for the F4 offspring at P360 in the control vehicle group (CO; A, D, and G) and DEHP group in the double-cross line (DC; B, C, E, F, H, and, I) are shown. The stages of seminiferous epithelial cycle are noted in control sections (A, D, and G). Arrowheads indicate areas where small vacuoles were observed. Arrows indicate areas where large vacuoles were observed. Boxed areas are magnified in b‘ and f‘, showing multinucleated cells. Hematoxylin and eosin-stained epididymal cauda cross sections for the F4 offspring at P360 in the CO group (J) and DEHP group (K and L). Bar = 50 μm. Photomicrographs are representative of n ≥ 3 offspring.

To compare the frequency of testicular abnormalities, the percentages of moderate to severe abnormal seminiferous tubules were determined for the DEHP-treated and vehicle-treated groups. The testes from F1 offspring at P120 of the DEHP-exposed group had a statistically significant increase of 5.0-fold in the percentage of abnormal tubules compared to the testes from the vehicle control group (Fig. 6A). In F2 and F3 offspring at P120, the testes from the three breeding lines of the DEHP-exposed group were found to have significant changes in the percentage of abnormal tubules compared to their vehicle counterparts (Fig. 6, B and C), although the range of fold changes (2.8- to 3.2-fold) was lower than the 5.0-fold change observed for testes from F1 offspring.

FIG. 6

Fold changes in abnormal tubules for F1–F3 offspring at P120 and F4 offspring at P360. Moderately and severely disorganized seminiferous tubules were counted and the percentage abnormality calculated by counting the total number of tubules on three or four sections spaced 50 μm apart per testis from n ≥ 3 offspring (≥9–12 sections). The fold change for the DEHP group was calculated by the percentage average of abnormal tubules in the control vehicle group (CO) set to 1. The fold changes in abnormal tubules were graphed for the F1 offspring at P120 (A); for the F2 and F3 offspring at P120 from the maternal (M), paternal (P), and double-cross (DC; B and C) lines; and for the F4 offspring at P360 from DC (D). Fold change values are means ± SEM. Pooled t-tests were conducted for the control and DEHP values; ***P < 0.001, **P < 0.01, *P < 0.05, no asterisk indicates P > 0.05.

In F4 offspring at P360, we found a 6.3-fold increase in abnormal tubules from the DEHP group compared to the vehicle control (Fig. 6D), which is more than the 2.8- to 3.2-fold increases observed for F2 and F3 offspring at P120, suggesting that testicular germ cell disorganization had increased with aging. F4 offspring at P360 had 66.6% of their tubules with moderate to severe testicular abnormalities compared to 27.1% of tubules in F2 and F3 offspring at P120. These results clearly show that the DEHP exposure of F0 dams induced testicular abnormalities in offspring of multiple generations (F1–F4), and that the frequency of testicular germ cell disorganization and abnormalities was higher in F1 compared to F2 and F3 generations and the frequency and severity increased with aging.

Germ Cell Apoptosis Did Not Contribute to F3 Offspring Phenotypes at P120

To determine if germ cell apoptosis contributed to the germ cell loss and germ cell disorganization in F1–F3 offspring at P120, TUNEL analyses were performed on the testes derived from DEHP- and corn oil vehicle-treated groups for all three breeding lines. The majority of the TUNEL-positive cells in the control group were located near the basement membrane (Fig. 7A), whereas the TUNEL-positive cells in the DEHP group were located near the basement membrane and also one layer above (Fig. 7, B and C). There were also tubules containing sloughed germ cells that did not have any positive stain (Fig. 7C, arrow). The apoptotic tubules with four or more TUNEL-positive cells were counted. There were no statistically significant differences at P < 0.05 for the F1 offspring from the DEHP-treated group (Fig. 7D) or the F2 and F3 offspring from the DEHP-treated group of all three breeding lines (Fig. 7, E and F). Thus, for DEHP treatment of F0 dams between E7 and E14 in mice, the germ cell disorganization observed in the F1, F2, and F3 offspring at P120 was independent of germ cell apoptosis. However, we cannot rule out that apoptosis did not occur earlier in the neonatal developmental window. Systematic developmental studies are necessary to determine if apoptosis contributed to DEHP-affected phenotypes.

FIG. 7

Fold changes in apoptotic tubules in F1–F3 offspring at P120. TUNEL assay was performed on three or four sections spaced 50 μm apart per testis from n ≥ 3 offspring (≥9–12 sections). Tubules with four or more apoptotic cells and the total number of tubules were counted to calculate the percentage apoptosis. Examples of tubules with apoptotic cells for the control (A) and DEHP (B and C) groups are shown. Arrow in C indicates abnormal seminiferous tubule with sloughed germ cells and no TUNEL-positive cells. The fold change for DEHP group was calculated by the percentage average of apoptotic tubules in the control vehicle group (CO) set to 1. The fold changes in apoptotic tubules were graphed for the F1 offspring at P120 (D) and for the F2 and F3 offspring at P120 from the maternal (M), paternal (P), and double-cross (DC; E and F) lines. Fold change values are means ± SEM. Pooled t-tests were conducted for the control and DEHP values and there were no significant differences at P < 0.05. Bars in A–C = 100 μm.

DEHP Decreased Sperm Count and Motility in F2 and F3 Offspring

To determine whether sperm count and motility changed transgenerationally, epididymal sperm from the F1, F2, F3, and F4 offspring at P120 of DEHP- and vehicle-treated groups were analyzed. There was a significant decrease in sperm count, on average 7.8% for F2, F3, and F4 offspring in all three lines (Fig. 8, B–D), except for the decrease in the sperm count for the maternal line for the F2 offspring, which was not significant at P < 0.05, but was close to significant (P = 0.0594). The F1 generation sperm counts had a decrease of 27.0%, more than 3-fold higher than those of the F2–F4 generations (Fig. 8A). This is not unexpected, because both germ cells and somatic cells of F1 embryos were directly exposed to DEHP. In terms of sperm motility, sperm from F1–F4 offspring of vehicle-treated F0 dams had a forward progressive motility at a frequency of 78.7% on average, whereas sperm from F1–F4 offspring of DEHP-treated F0 dams exhibited a forward progressive motility at a frequency of 53.0% on average, a decrease in motility compared to controls.

FIG. 8

Epididymal sperm count from F1–F4 offspring at P120. Epididymal sperm counts were performed on caudal sperm from n ≥ 6 offspring for the F1, F2, F3, and F4 generations. The percentage change in sperm count for DEHP group was calculated against the control group (CO) sperm count set to 100%. The percentage sperm count were graphed for the F1 (A), F2 (B), F3 (C) and F4 (D) offspring at P120 from the maternal (M), paternal (P), and double-cross (DC) lines. The percentage age values are means ± SEM. Pooled t-tests were conducted for the control and DEHP values; ***P < 0.001, **P < 0.01, *P < 0.05, no asterisk indicates P > 0.05.

Stem Cell Function of F3 Offspring Is Impaired

Recent reports that phthalates decreased the proliferation of the SSC line [41, 42] led to the investigation of SSC function. Germ cells from F3 offspring (P24 to P31) of the DEHP-treated group in the double-cross line and nontreated CD1 control mice were used as SSC donors and transplanted into W/W^V recipient mice, which have no endogenous germ cells undergoing meiosis. Germ cells from F3 offspring were chosen for studying the transgenerational effects because they were not directly exposed to DEHP, whereas the germ cells in F2 offspring were derived from the primordial germ cells in the F1 offspring that were directly exposed to DEHP. The weights of testes from P30 animals in the control and DEHP groups from the F3 generation were not significantly different at P < 0.05 (data not shown).

The hematoxylin and eosin-stained testis cross sections from the W/W^V recipients injected with nontreated control CD1 germ cells revealed the presence of seminiferous tubules containing germ cells (Fig. 9A), a 65.1% donor germ cell-derived recovery (Fig. 9J). In contrast, the testis cross sections of W/W^V recipients injected with germ cells from F3 offspring of the DEHP-treated group displayed a dramatic decrease in tubules containing donor-derived germ cells (Fig. 9B, arrows), a 4.7% recovery (7.3% of the control) (Fig. 9J). Anti-DDX4 antibody immunostaining (examples shown in Fig. 9, D–F) and anti-γH2AX antibody immunostaining (examples shown in Fig. 9, G–I) and counting (Fig. 9, K and L) confirmed that there are fewer tubules with meiotic and postmeiotic germ cells. DDX4 is found in spermatogonia and spermatocytes, as well as in spermatids, staining more than two layers of germ cells [43]; γH2AX is localized to preleptotene, leptotene, zygotene, and early pachytene spermatocytes in the testis [44]. As an endogenous negative control, noninjected W/W^V testes were used. Noninjected W/W^V testes did not contain any donor-derived germ cells (Fig. 9C) and anti-γH2AX (Fig. 9I) immunostains did not reveal any immune-positive cells.

FIG. 9

W/W^V recipient mice injected with donor germ cells from F3 offspring of DEHP-treated F0 dams. Testicular cross sections from W/W^V recipients injected with germ cells from untreated CD1 control mice (A, D, and G), germ cells from F3 offspring derived from DEHP-exposed F0 dams (B, E, and H), and a noninjected W/W^V control testis (C, F, and I) were stained with hematoxylin and eosin (A–C) or immunostained using anti-DDX4 antibody (D–F) or anti-γH2AX antibody (G–I). To quantify the extent of recovery, tubules containing donor-derived germ cells (J), tubules with 2 or more layers of anti-DDX4 immunostain (K), tubules with anti-γH2AX immunostain (L), tubules with round spermatids (M), and tubules with elongated spermatids (N) were counted and graphed. Values are means ± SEM for n ≥ 3 offspring and for two to six sections separated by 50 μm per testis. Pooled t-tests were conducted for the control and DEHP values; ***P < 0.001, **P < 0.01. Bars = 400 μm (A–F) and 50 μm (G–I). The negative controls showed no staining with only secondary antibody present, but without γH2AX or DDX4 primary antibodies (data not shown).

To further examine if the recovered tubules had a complete spermatogenesis, the recovered tubules were counted for the presence of round or elongated spermatids. In the W/W^V recipients injected with control CD1 germ cells, 53.3% and 55.6% of the tubules contained round and elongated spermatids, respectively (Fig. 9, M and N). In contrast, there was a dramatic decrease in round and elongated spermatids in the W/W^V recipients injected with F3 offspring germ cells from the DEHP-treated group, with only 4.5% (8.4% of the control) of the tubules containing round spermatids and 1.8% (3.2% of the control) of the tubules containing elongated spermatids (Fig. 9, M and N).

Stem Cells from F3 Offspring of the DEHP-Treated Group Have Decreased Colonization

To determine if there was any difference in the number of undifferentiated spermatogonia in the W/W^V recipient testes, anti-DMRT1 antibody was used. Anti-DMRT1 strongly immunostains undifferentiated spermatogonia (white arrows in Fig. 10, A, B, and C) and lightly stains differentiated spermatogonia and Sertoli cells [45]. Tubules containing the brightest-staining undifferentiated spermatogonia were counted in the W/W^V recipients injected with control germ cells, in F3 germ cells from the DEHP group, and in the noninjected W/W^V testes. The W/W^V control recipients injected with untreated CD1 donor cells had 58.6% of tubules with undifferentiated spermatogonia (Fig. 10I), whereas the W/W^V recipients injected with F3 DEHP donor cells had 12.2% of tubules with undifferentiated spermatogonia (20.8% of the control). Thus, these results indicate that a large decrease (∼79%) in stem cell colonization compared to the controls.

FIG. 10

W/W^V recipient testicular tubules containing undifferentiated, differentiated, and premeiotic germ cells. Immunofluorescence and immunohistochemistry were performed using an anti-DMRT1 antibody (A–D) and an anti-STRA8 antibody (E–H), respectively, on testicular sections from W/W^V mice injected with germ cells from untreated control CD1 mice (A and E) and germ cells from F3 offspring of DEHP-treated F0 dams (B and F), and noninjected W/W^V control mice (C and G). Anti-DMRT1 antibody immunostains undifferentiated spermatogonia strongly (examples shown by brightly fluorescent cells marked by white arrowheads in A, B, and C). Using Adobe Photoshop CS (Adobe Systems Inc., San Jose, CA) the Alexa-555 color (red) was postprocessed to give a green color for better visualization. Anti-STRA8 antibody immunostains differentiated spermatogonia and premeiotic spermatocytes (examples indicated by brown stained cell nuclei in E, F, and G). Negative staining of the secondary antibody (negative controls) only on testis sections for anti-DMRT1 antibody and anti-STRA8 antibody did not show any staining (D and H). Bars = 50 μm (A–D) and 20 μm (E–H). Photomicrographs are representative of n ≥ 3 animals. To quantify, DMRT1-positive tubules containing brightly stained undifferentiated spermatogonia (I) and STRA8-positive tubules containing differentiated spermatogonia and premeiotic spermatocytes (J) were counted as well as the total tubules. Values are percentage means ± SEM from n ≥ 3 animals and two to six sections per testis spaced apart by 50 μm. Pooled t-tests were conducted for the control and DEHP values; ***P < 0.001, *P < 0.05 (I and J).

The recipient testes were further examined for tubules that contained differentiating spermatogonia and early meiotic spermatocytes. The antibody against stimulated by retinoic acid gene 8 (STRA8) was chosen to examine cell types (represented by darkly staining brown nuclei), differentiating spermatogonia (A1–A4, Int, and B) and early meiotic spermatocytes (preleptotene and leptotene) [46]. The W/W^V control recipients injected with untreated CD1 donor cells had 46.3% of tubules with differentiated spermatogonia and early meiotic spermatocytes expressing STRA8, whereas 8.9% of the tubules (19.5% of the control) in the recipient testes injected with F3 germ cells from the DEHP-treated group had STRA8-positive cells (Fig. 10J). Together, these results indicate that there was no further decrease in donor-derived spermatogenic cells after the initial large decrease in stem cell colonization and proliferation observed with the DMRT1 immunostaining. However, there was a second large drop in donor-derived spermatogenic cells from the beginning of meiosis (19.5% STRA8-positive cells) to the round spermatid stage (Fig. 9, 8.4% of control tubules with round spermatids).

Germ Cell Disorganization Is an Inherent Property of F3 Stem Cells from DEHP-Treated Group

Interestingly, when the tubules from the W/W^V recipients transplanted with the F3 offspring germ cells were examined, they had germ cell disorganization (Fig. 11, B, C, E, F, H, and I) very similar to the germ cell disorganization observed in testes from the F3 and F4 offspring of the DEHP-treated group at P120 and P360 (Figs. 4 and 5). The tubules appeared to lack layers of germ cells, were disorganized, and had areas where vacuoles were observed (Fig. 11B) or contained multinucleated cells (Fig. 11, C, F, and I, arrows). On the other hand, the tubules from the recipients injected with untreated CD1 germ cells had normal morphology with readily observable normal stages of seminiferous epithelium (Fig. 11, A, D, and G).

FIG. 11

Morphology of recovered tubules in W/W^V mice injected with germ cells from F3 offspring of DEHP-exposed F0 dams. Testicular cross sections from W/W^V recipients injected with germ cells from untreated CD1 control mice (A, D, and G), showing normal stages of seminiferous epithelium, and germ cells from F3 offspring from DEHP-exposed F0 dams (B, C, E, F, H, and I), showing abnormal testicular morphology such that the stages of the seminiferous epithelium could not be assigned. Arrows indicate multinucleated cells. Bar = 50 μm. Photomicrographs are representative of n ≥ 3 animals.