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BOR - Papers in Press, published online ahead of print September 22, 2004.
Biol Reprod 2004, 10.1095/biolreprod.104.033282
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BIOLOGY OF REPRODUCTION 72, 451–459 (2005)
DOI: 10.1095/biolreprod.104.033282
© 2005 by the Society for the Study of Reproduction, Inc.

Birth of Pouch Young after Artificial Insemination in the Tammar Wallaby (Macropus eugenii)1

Damien B.B.P. Paris3, David A. Taggart3,4, Geoff Shaw3, Peter D. Temple-Smith3, and Marilyn B. Renfree2,3

Department of Zoology,3 University of Melbourne, Victoria 3010, Australia Royal Zoological Society of South Australia,4 c/o School of Earth and Environmental Science, University of Adelaide, South Australia 5005, Australia


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Timing of artificial insemination (AI) in marsupials is critical because fertilization must occur before mucin coats the oocyte during passage through the oviduct. In this study, timing and the site of insemination were examined to develop AI in the tammar wallaby (Macropus eugenii). Birth and postpartum (p.p.) estrus was synchronized in 46 females. Epididymal spermatozoa (n = 4) or semen collected by electroejaculation (n = 42) were inseminated early (4–21 h p.p.) into the urogenital sinus (n = 7), the anterior vaginal culs de sac (n = 7), the uterus by transcervical catheter (n = 5), or the uterus by injection (intrauterine artificial insemination, IUAI) (n = 5). A further 16 females were inseminated late (19–48 h p.p.) by IUAI. All females were monitored for birth. A third group of six females was inseminated late (21–54 h p.p.) by IUAI and 0.4–6.6 h later, sperm had reached the oviduct in all animals. In total, an oocyte to which spermatozoa were attached was recovered and two young were born after IUAI using epididymal (n = 1) or electroejaculated (n = 2) spermatozoa, but no young resulted from insemination at other sites. Two females were successfully inseminated at 43 and 47 h p.p., later than most other animals, and the third was inseminated much earlier (18 h p.p.) but with highly motile spermatozoa. These young represent the first macropodids born by AI and the first marsupials conceived using epididymal spermatozoa.

assisted reproductive technology, female reproductive tract, fertilization, marsupial, ovulation, sperm motility and transport


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Artificial insemination (AI) has been used increasingly in wild species [1] but has only been attempted in three marsupials: the tammar wallaby (Macropus eugenii) [2, 3], the brushtail possum (Trichosurus vulpecula) [2, 4, 5], and the koala (Phascolarctos cinereus) [6]. The only confirmed births after AI in marsupials have been in the koala, which appears to be an induced ovulator. Nonsurgical AI into the urogenital sinus in the koala has a high (37%) success rate [6]. However, in most other marsupials, ovulation is spontaneous [7] and fertilization must occur shortly after ovulation because the oocyte is transported through the oviduct in less than 24 h, during which it acquires a mucin layer that inhibits sperm penetration [8–10] and a tertiary egg membrane, the shell coat [11]. Insemination in marsupials must therefore be timed so that spermatozoa can reach the oocyte in the ampulla of the oviduct before deposition of the mucin and shell coat layers.

In an attempt to regulate the timing of ovulation for AI in the tammar wallaby and brushtail possum, various superovulation protocols have been developed [2, 5]. In the possum, one third of ovulated oocytes were fertilized after intrauterine artificial insemination (IUAI) 0–6 h before and 2–18 h after expected ovulation, using varied doses of eCG/ GnRH as well as eCG/LH [2]. A fertilization rate of 42% was reported in the brushtail possum after IUAI 1–4 h before eCG/LH-induced ovulation and 1.5–2.5 h before FSH/ LH-induced ovulation [5]. Vaginal insemination 16–17 h before FSH/LH-induced ovulation resulted in a 40% fertilization rate in this species [5]. In FSH/LH-treated tammar wallabies, the fertilization rate was 45% and 70% after IUAI 4–6 h and 5–7 h, respectively, before the expected ovulation [2, 3]. From these data, the optimal time of IUAI in both the brushtail possum and tammar wallaby appears to be less than 7 h before artificially induced ovulation.

Surprisingly, despite these high rates of fertilization, no live births have been reported in either species in which superovulation has been used for synchronization. Embryos produced by AI after superovulation in the monovular tammar and possum have a high rate of degeneration [2]. It is possible that the uterine environment of superovulated monovular marsupials may not be able to sustain pregnancies to term [12]. Further, superovulated oocytes in the possum have improperly formed tertiary mucin and shell layers [13] that may be detrimental during cleavage stages and blastocyst expansion [11]. Thus, alternative methods of synchronization that do not rely on exogenous hormone administration should be explored.

The tammar wallaby has a postpartum (p.p.) estrus and mates within 1–6 h of birth (1.3 ± 0.8 h) [14]. Time of ovulation is variable, but it usually occurs between 43 and 60 h p.p. [10] after which the fertilized egg develops to the blastocyst stage and then enters diapause in lactating females [7]. Birth can be synchronized by removing pouch young (RPY) attached to the teat, which results in reactivation of the blastocyst and birth 26.4 ± 0.2 days later [15]. Thus, RPY and birth can be used to synchronize and time estrus and ovulation independent of exogenous gonadotrophins.

In the tammar, after natural mating, sperm reach the cervix within 40 min of copulation, the uterus by 4 h, and are in the lower oviduct within 6 h. Spermatozoa are still present in the uterus and lower oviduct 24 h postcoitum (p.c.) [16]. In addition, IUAI in superovulated tammars 4–7 h before induced ovulation results in up to 7 fertilized oocytes per female [2, 3]. In natural cycles, because ovulation occurs 43–60 h p.p., birth can be used to time insemination of spermatozoa into different sites of the female reproductive tract relative to an expected ovulation and when spermatozoa are found in these regions after natural mating.

The aim of this study was to test different sperm preparations, insemination sites, and insemination timing to achieve successful AI in the tammar wallaby.


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

Tammar wallabies from Kangaroo Island, South Australia, were held in a breeding colony in Melbourne in large outdoor enclosures. Pasture was supplemented with lucerne hay cubes, fresh vegetables, and water ad libitum. A total of 46 females of proven fertility and 21 adult males were used. Sperm from single ejaculates from five males and from epididymal aspiration of two males were used to inseminate one female each. Sperm from single ejaculates from another seven males and from dissected epididymides of one male were split to inseminate 2–4 females each, while ejaculates collected on 2–3 separate occasions from a further six males were used to inseminate 1–2 females each. Anesthesia was induced with an intramuscular dose of tiletamine and zolazepam (50 mg Zoletil 100; Virbac Pty. Ltd., Peakhurst, NSW, Australia) and maintained with 2–5% isofluorane (Abbott Australasia Pty. Ltd., Kurnell, NSW, Australia) in 1 L/min oxygen. All experiments were approved by the Animal Experimentation Ethics Committees of the University of Melbourne.

Criteria for Estimating the Time of Birth

Birth in 12 females was either observed directly or detected shortly thereafter based on the presence of a pouch young not yet on the teat [15], and the urogenital opening was examined for signs of blood or fetal membranes. The skin color of the pouch young was recorded at birth and, in six females, at one or two additional times from 0.5 to 20 h p.p., and in three females, every 3–4 h during daylight from 3 to 11 h p.p., 13–24 h p.p., and 27–55 h p.p., respectively. A set of criteria to estimate time of birth was developed (Fig. 1).



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  		FIG. 1. Skin color change over time in tammar wallaby pouch young (PY) after birth: (a) fiery-red PY with very red prominent blood vessels and not on teat < 1 min p.p., (b) fiery-red PY with faint blood vessels and on teat 1.8 h p.p., (c) dull-red PY 14.4 h p.p., (d) pink PY < 23.6 h p.p., and (e) timeline of color change. Figure 1a reproduced from [19] by permission of Surrey Beatty and Sons and Dr. D. Parer

Synchronization of Estrus

Females used for AI were isolated from males and pregnancy was synchronized by the removal of RPY [15, 17]. From the morning of Day 26, females were checked for the presence of pouch young approximately every 12 h to minimize disturbance to the animals. The time the pouch young was first detected, its color and the condition of the female's urogenital opening were recorded. The time of birth was estimated using the neonate color and female urogenital appearance criteria developed above (see Fig. 1 and Results). Where the time of birth estimated using these criteria was earlier than the previous check, birth was estimated to have occurred 30 min after the previous checking time to allow for animal release and return to normal activity.

Collection of Blood Samples

Blood (10 ml) was collected using syringes coated with saline containing 125 IU/ml heparin (David Bull Laboratories, Mulgrave, Victoria, Australia) or K3-EDTA coated vacuette blood tubes (Greiner Bio-One, Kremsmuenster, Austria). Samples were collected from the lateral tail vein within 5 min of administration of anesthetic and before semen collection in males or AI in females and again at the time of death in those females where tracts were removed. Female plasma for hormone analysis and a 1 ml leukocyte-rich fraction (buffy coat) for DNA extraction were stored at –20°C. Male whole blood or buffy coats for DNA extraction were also stored at –20°C.

Semen Collection, Processing, and Assessment

Collection by electroejaculation Forty-two semen samples were obtained under anesthesia by electroejaculation (modified from [18]). Semen was collected using a 10-mm-diameter rectal probe with three longitudinally oriented, surface-mounted 30-mm strip electrodes connected to a 20-Hz sine-wave generator (CGS Products Pty. Ltd., Trafalgar, Victoria, Australia). The penis was cleaned with swabs moistened with 0.9% saline and placed into a sterile 14-ml round-bottom tube (Becton Dickinson Labware, Franklin Lakes, NJ) containing collection medium. The probe was lubricated (K-Y Jelly; Johnson and Johnson, Botany, NSW, Australia) and inserted approximately 10 cm into the rectum with the electrodes ventrally oriented in close apposition to the prostate. The electroejaculation sequence included three series separated by 30-sec rest periods. Each series consisted of ten 3-sec stimuli at each of three voltages. Series 1 and 2 were conducted at 2, 3, and 4 V and series 3 at 3, 4, and 5 V.

Collection of epididymal spermatozoa Four samples were obtained under anesthesia from the cauda epididymidis by aspiration (n = 2) or whole caudal dissection (n = 2). For epididymal aspiration, a 26-gauge needle was used to puncture the scrotal sac into the caudal epididymidis, and 0.1 ml medium flushed in and out until it appeared cloudy or milky. For caudal dissection, the epididymis was dissected out and the caudal epididymidis excised and sliced finely in 1.5 ml of collection medium.

Collection media Semen was collected directly into known volumes of prewarmed (35°C) culture medium. Dulbecco modified Eagle medium containing 20 mM HEPES and 4.5 g/L glucose (D-MEM-) (Thermo Trace Ltd., Noble Park, Victoria, Australia); D-MEM- supplemented with 4% fetal bovine serum (FBS) and 50 IU/ml, 50 µg/ml penicillin, streptomycin (Pen/Strep) (both; Thermo Trace Ltd., Noble Park, Victoria, Australia) (D-MEM+); or Hams F-10 medium containing 25 mM HEPES and 146 mg/ L L-glutamine (Invitrogen Corp., Mulgrave, Victoria, Australia) supplemented with 5% FBS and 50 IU/ml, 50 µg/ml Pen/Strep (Hams F-10+) was used. Preliminary assessment indicated no apparent difference in sperm motility in each medium. Samples collected in D-MEM+ were resuspended in Hams F-10+ before AI.

Assessment and processing of spermatozoa For each semen sample, except the four epididymal sperm samples, ejaculate volume was measured and coagulation properties were scored on a 0–3 scale, where 0 = no coagulation and 3 = solid plug formation. Samples with large ejaculate volume were allocated to the urogenital sinus treatment group (refer below). Samples were maintained at 35°C and left for 10 min to allow spermatozoa to migrate from the plug or sliced caudal tissue into the medium. The sperm-enriched medium was then assessed for sperm concentration, percentage motile spermatozoa, motility rating, and sperm motility index (modified from [19]). Motility rating is a subjective assessment of speed and directionality of the largest proportion of spermatozoa in a given sample and was graded on a 0–4 scale, where 0 = none, 1 = nonprogressive motility, 2 = slow forward progression, 3 = intermediate forward progression, and 4 = rapid straight-forward progression. Sperm motility index is a motility estimate for the entire population of spermatozoa in a given sample and was measured on a scale of 0–400, where 0 = 100% of spermatozoa are nonmotile and 400 = 100% of spermatozoa have grade-4 motility. To calculate, sperm motility index = (0 x percentage of spermatozoa of motility rating 0) + (1 x percentage of spermatozoa of motility rating 1) + (2 x percentage of spermatozoa of motility rating 2) + (3 x percentage of spermatozoa of motility rating 3) + (4 x percentage of spermatozoa of motility rating 4). If sperm concentration was high, an aliquot of the sperm-rich medium was used directly for insemination in at least one or sometimes multiple females. Otherwise, samples were spun 5 min at 250 x g and the pellet resuspended in a smaller volume of fresh medium. The total number of spermatozoa inseminated was then calculated from the final insemination concentration.

Artificial Insemination Procedures

Females were inseminated under anesthesia between 4 and 54 h after the estimated time of birth. Semen was deposited early (4–21 h p.p.) into four different regions of the reproductive tract of 24 females (see below). Another 22 were inseminated late (19–54 h p.p.) into the uterus, 6 of which were killed at specific times and the location of spermatozoa in the reproductive tracts recorded (see below). All other females were monitored for birth 26–35 days later. A tail tip was collected from all offspring born and stored at –20°C for DNA extraction and microsatellite paternity analysis.

Insemination into the urogenital sinus Semen was inseminated into the urogenital sinus of seven females using a modified sterile 10-ml yellow screw-cap tube (Sarstedt Australia, Pty. Ltd., Technology Park, SA, Australia). The tip of the tube containing the semen was cut off to create a wide-bore hole and a plunger from a 5-ml syringe (Terumo Medical Corp., Elkington, MD) was used to inject the whole ejaculate in medium through the tube into the urogenital opening.

Insemination into the anterior vaginal culs de sac Semen was inseminated into the anterior vaginal culs de sac of seven females using a 5-French 30-cm silicon balloon HSG catheter (Cook IVF, Eight Mile Plains, Queensland, Australia) introduced at the urogenital opening and navigated through the median vagina until the tip was adjacent to the cervices. The balloon was inflated with 0.4 ml sterile 0.9% saline to seal the median vagina and 1.5–1.9 ml sperm-rich medium injected. The catheter was held in place for 5 min to prevent loss of spermatozoa down the median vagina.

Insemination through the cervix Semen was inseminated transcervically into the uterus of five females at laparotomy [20]. The side of impending ovulation and side of parturition were determined at laparotomy. The cervices were exposed by an incision in the anterior vaginal culs de sac using sterile procedures. Sperm-rich medium (0.2 ml) was delivered through the cervix into the uterus ipsilateral to the Graafian follicle via a 1.5-mm diameter medical-grade catheter tube (Dural Plastics and Engineering, Auburn, NSW, Australia) attached to a 21-gauge needle and 1-ml syringe (Terumo Medical Corp.). Postoperatively, animals were given prophylactic antibiotic: 1 ml intramuscular dose of procain penicillin (150 mg) and benzathine (115 mg) (Duplocillin: Intervet Australia Pty. Ltd., Toongabbie, NSW, Australia).

Intrauterine insemination: i) early IUAI (11–18 h p.p) Semen was inseminated directly into the uterus of five females (IUAI). Only the nonparturient uterus ipsilateral to the Graafian follicle was injected with 0.2 ml sperm-rich medium through the uterine wall into the lumen using a 26-gauge needle and 1-ml syringe (Terumo Medical Corp.). The uterus was seen to swell upon injection. The incision was closed and antibiotics administered as described above.

Intrauterine insemination: ii) late IUAI (19–48 h p.p) Sperm-rich medium (0.1–0.2 ml) was delivered by IUAI and laparotomy into 16 females as described above. Seven of these females were inseminated in mid-June, when females normally become seasonally quiescent. To synchronize births as far as possible, these seven animals were injected with 20 mg bromocriptine (Novartis, Basel, Switzerland) 8 days after insemination [21].

Intrauterine insemination: iii) late IUAI (21–54 h p.p.) with tracts flushed Six females were inseminated with 0.15–0.2 ml sperm-rich medium into the uterus as described above. Animals were killed, reproductive tracts removed at 0.4, 1.2, 6.0, 6.0, 6.4, and 6.6 h post-IUAI and ligated into 13 major anatomical sections: urogenital sinus, two posterior lateral vaginae, median vagina, two anterior lateral vaginae, cervices and anterior vaginal culs de sac, two uteri, two lower oviducts, and two upper oviducts. Spermatozoa and embryos were recovered by flushing with sterile 0.9% saline. Sperm numbers were counted for each section using a hemocytometer [19] and follicle size was measured with vernier calipers.

Progesterone and Estrogen Assays

The assay for progesterone was based on the method of Renfree et al. [22] validated for tammar plasma using antiserum C-9817 (Bioquest Ltd., North Ryde, NSW, Australia) modified as follows: 750 µl of female plasma was extracted, antiserum was used at 1:60 000 final working dilution, and 2 ml of scintillation fluid (Ultima Gold: Packard Bioscience Co., Meriden, CT) was used. Samples were assayed in two batches. The mean extraction efficiencies were 75.4% and 80.5%, respectively. Assay sensitivity was 44.2 pg/ml (10 pg/tube) for assay 1 and 70.4 pg/ml (17 pg/tube) for assay 2. The intraassay coefficient of variation (CV) for two quality-control pools containing 916.7 ± 25.4 (mean ± SEM, n = 5) and 1241.0 ± 39.2 pg/ml progesterone (n = 8) was 6.2% and 8.9%, respectively. The interassay coefficient of variation for quality-control pools containing 1070.4 ± 44.6 (n = 7) and 907.0 ± 43.9 pg/ml progesterone (n = 6) was 11.0% and 11.8% respectively. Solvent and buffer blanks were below the sensitivity of the assay.

The assay protocol for estradiol was based on the method of Shaw and Renfree [23] validated for tammar plasma using antiserum C-6181 (Bioquest Ltd.) modified as follows: 800 µl of female plasma was diluted with 800 µl distilled water and heated at 92–95°C to denature binding proteins in plasma; ether extracts were dried and redissolved in 500 µl assay buffer; estradiol-17ß standards ranged from 2 to 800 pg, and 2 ml of scintillation fluid (Ultima Gold: Packard Bioscience Co.) was used. Samples were assayed in two batches. The mean extraction efficiencies were 81.0% and 83.0%, respectively. Assay sensitivity was 12.7 pg/ml (3.3 pg/tube) for assay 1 and 7.5 pg/ml (<2 pg/tube) for assay 2. The intraassay CVs for two quality-control pools containing 29.8 ± 1.1 (mean ± SEM, n = 6) and 30.8 ± 1.0 pg/ml estradiol (n = 6) were 8.9% and 7.8%, respectively. The interassay CVs for quality-control pools containing 26.95 ± 2.6 (n = 5) and 37.4 ± 3.3 pg/ml estradiol (n = 3) were 21.9% and 15.3%, respectively. Solvent and buffer blanks were 4 and 4.7 pg/tube, respectively, in assay 1 and 9.5 pg/tube and below the sensitivity of the assay, respectively, in assay 2.

Microsatellite Paternity Analysis

DNA was extracted using the Qiagen DNeasy Tissue Kit (Qiagen Pty. Ltd., Clifton Hill, Victoria, Australia) and methods outlined in the DNeasy Tissue Handbook (May 2002). Blood and buffy coat samples (100 µl) were processed according to the ‘Isolation of DNA from whole non-nucleated blood’ protocol and pouch young tails (6 mm) were extracted using the ‘DNeasy protocol for rodent tails,’ both modified by performing two 100-µl elutions in AE buffer (Qiagen) instead of one 200-µl elution, to improve DNA yield. The concentration of extracted DNA was determined by loading 1- and 2-µl aliquots of sample into an 0.8% agarose gel (Scientifix, Cheltenham, Victoria, Australia) stained with 0.8 µg/ml ethidium bromide (Sigma, Castle Hill, NSW, Australia) and electrophoresed at 110 V for 30 min. DNA of known concentration (12.5, 25, 50, 75, 100, and 150 ng) was also loaded and visually compared with unknown samples. DNA samples were diluted to 2.5 ng/µl and analyzed at the Australian Genome Research Facility (AGRF, Parkville, Victoria, Australia) by PCR amplification on a Peltier Thermal Cycler 225 (MJ Research Inc., Waltham, MA) and gel separation and allele sizing using an ABI PRISM 377 automated DNA sequencer (Applied Biosystems, Foster City, CA).

Paternity was determined using five polymorphic microsatellite loci (Me1, Me14, Me15, Me16, and Me17) specific for the tammar wallaby [24]. The 5' end of the forward primer in each primer pair was end labeled with FAM or TET ABI dyes (Sigma). PCR reactions contained 1x PCR reaction buffer (Mg2+ free), {chi} mM MgCl2 (where {chi} = 1.5 for Me 1, 14, 16, 17 and {chi} = 2.0 for Me 15), 0.2 mM dNTPs, 1.2 U Taq DNA polymerase 1 (all from Invitrogen Corp.), 0.5 µM each primer, 5 ng genomic DNA, and made up to 20 µl total reaction volume using autoclaved milli-Q dH2O. PCR cycling conditions consisted of an initial denaturation step of 94°C for 5 min, then 40 cycles of (94°C for 45 sec, annealing temperature [TA°C] for 30 sec, 72°C for 45 sec), and finally 72°C for 10 min. Annealing temperatures (TA) were 56, 52, 60, 60, and 56°C for Me1, 14, 15, 16, and 17 primers, respectively.

Although females were isolated from males before and after AI at the time of estrus, on rare occasions, males can jump dividing fences to reach females. Paternity analysis was used to confirm that each sperm donor was the father of AI offspring. Paternity was assigned by determining the allele frequency in the wallaby colony and analyzing parentage based on known mothers using Cervus 2.0 software [25].

Statistical Analysis

Values are presented as mean ± SEM. Raw data were analyzed for normal distribution, equal variance between groups, and the presence of outliers [26]. Data for semen volume, sperm concentration, total number of spermatozoa, and progesterone concentration were log10 transformed and percentage motile spermatozoa and motility rating were reflected, then log10 transformed to normalize the sample data [26]. Semen parameters were grouped by AI treatment and differences in the mean values were compared by unbalanced ANOVA and post hoc pairwise comparison of means with a Tukey test using SYSTAT 9.01 software. Relationships between the time of insemination and progesterone or estradiol concentration were tested by Pearson correlations and significance determined by Bonferroni adjusted probabilities using SYSTAT 9.01 software. P < 0.05 were considered to be significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Changes in the Appearance of Pouch Young after Birth

Birth was directly observed in 7 females. By 5 min after birth, the pouch young were attached to the teat, and after 40 min, blood could no longer be detected at the urogenital opening of the female (Fig. 1). Pouch young were fiery red between the time of birth and 7.0 h p.p. (n = 11 observations) and blood vessels initially appeared very red and prominent but became faintly visible thereafter (Fig. 1, a, b, and e). Dull-red-colored pouch young were observed between 6.0 and 19.8 h p.p. (n = 9 observations) (Fig. 1, c and e). Pouch young turned pink as early as 17.6 h p.p. and remained this color until the conclusion of the experiment at 54.6 h p.p. (n = 11 observations) (Fig. 1, d and e). There was approximately a 1-h overlap of color transition from fiery red to dull red and a 2 h overlap from dull red to pink for different pouch young (Fig. 1e). With the exception of pink pouch young (18 h p.p.), midpoint values were chosen as estimates of the time of birth for each distinct condition according to the following: i) fiery-red pouch young not yet on the teat (3 min p.p.); ii) fiery-red pouch young on the teat with blood on the female urogenital opening (23 min p.p.); iii) fiery-red pouch young on the teat with a clean female urogenital opening (4 h p.p.); iv) dull-red pouch young (13 h p.p.).

Site of Insemination and Outcomes

Successful insemination was only achieved when semen was deposited into the uterus by IUAI (Fig. 2 and Table 1). One early and one late IUAI insemination resulted in the birth of pouch young, and an oocyte to which spermatozoa were attached was recovered following flushing of a late IUAI-treated female. These three females represent only 13.6% of females treated by IUAI. The oocyte recovered from the lower oviduct had supernumerary spermatozoa (electroejaculated) distributed throughout its thin mucin layer as seen after natural mating [10] (Fig. 3a). The pouch young born in the early IUAI group was conceived using electroejaculated spermatozoa and is the first macropodid produced by AI (Fig. 3b), while the pouch young born in the late IUAI group is the first marsupial to be conceived using epididymal spermatozoa aspirated from the cauda (Fig. 3c).



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  		FIG. 2. Time of AI in 46 female tammar wallabies from six treatment groups. Black squares, birth of pouch young; black circle, sperm-covered oocyte; white circles, unsuccessful; white triangles, yet to ovulate. UGS, Urogenital sinus; AVC, anterior vaginal culs de sac; Trans, transcervical; IUAI, intrauterine artificial insemination


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  		TABLE 1. Semen parameters (mean ± SEM) of inseminates used in each group of female tammer wallabies and successful IUAI-treated individuals.a



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  		FIG. 3. IUAI success in the tammar wallaby: (a) oocyte enveloped with spermatozoa (arrow) flushed from the lower oviduct 6.4 h after insemination at 43 h p.p., (b) pouch young on the day of birth from a female inseminated 18 h p.p. using electroejaculated spermatozoa, (c) Day 18 pouch young from a female inseminated 47 h p.p. using epididymal spermatozoa. m, Mucoid coat; z, zona pellucida; p, perivitelline space; bar = 100 µm

Quality of Semen Used for Insemination

Semen quality differed significantly between the six treatment groups (P < 0.05) (Table 1). Ejaculates with large volumes were selectively used for urogenital sinus inseminations in an attempt to replicate the large quantities of semen deposited in the lower tract after natural mating. Ejaculates used in late IUAI were also significantly larger than those in the anterior vaginal culs de sac and had a significantly higher coagulation rating than both the anterior vaginal culs de sac and early IUAI inseminations (Table 1). The proportion of motile spermatozoa used in the urogenital sinus, anterior vaginal culs de sac, and transcervical inseminations were significantly higher than both late IUAI groups. Spermatozoa used to inseminate the urogenital sinus had a significantly lower motility rating and sperm motility index than the transcervical and anterior vaginal culs de sac groups as well as a significantly lower motility rating than the late IUAI group that was flushed. Among the IUAI groups, the concentration of spermatozoa in inseminates was significantly lower for early than late inseminations. With the exception of the late IUAI flushed group, females inseminated with the smallest volumes (0.2 ml) including transcervical, early and late IUAI, received significantly fewer spermatozoa (Table 1).

Spermatozoa used in the early successful insemination had the highest activity in terms of percentage motility, motility rating and sperm motility index than both late insemination successes (Table 1). For all three parameters, motility of this sperm sample was the highest among the five animals inseminated in this early IUAI group and higher than the mean values of all six groups of animals inseminated (Table 1). Among all successful attempts, the female inseminated late by IUAI and flushed had the highest sperm concentration and total number inseminated, while motility rating was also generally high. However, more than half of the animals treated received greater numbers of spermatozoa than this female, yet were unsuccessful (Table 1). The successful late IUAI attempt used semen of poor quality, with the fourth lowest percentage motility and sperm motility index as well as the sixth lowest number of spermatozoa compared with that used for all 46 females (Table 1).

Timing of Insemination

Ovulation occurs between 43 and 60 h p.p. in the tammar [10], and in our study, inseminations were conducted from 4–54 h p.p. (Fig. 2). No follicles had ovulated at the time of insemination up to 54 h p.p. in those animals assessed during surgical insemination (transcervical and IUAI groups). One female that later gave birth to a pouch young was inseminated at 47 h p.p. and a sperm-covered oocyte was recovered at 49.4 h p.p. from a second female after insemination at 43 h p.p. but sperm penetration was not assessed. These two animals were inseminated third and sixth latest among all 46 inseminations, respectively. Another female also gave birth to a pouch young after insemination much earlier, at 18 h p.p. However, 17 females inseminated between 18 and 47 h p.p., 2 of which occurred between 43 and 47 h p.p., failed to give birth (Fig. 2).

In five of the six females killed at 22, 34, 46, 49, and 60 h p.p., after insemination between 21 and 54 h p.p. (Fig. 2), Graafian follicles were 3.7, 4.2, 4.9, 4.2, and 4.0 mm, respectively. Only one of these six females had ovulated (by 49.4 h p.p.) and a sperm-covered oocyte was flushed from the lower oviduct (Figs. 2 and 3a). This animal was inseminated into the uterus at 43 h p.p. A second female flushed at 49 h p.p. had a Graafian follicle of 4.2 mm, while another small female still had not ovulated by 60 h p.p. and had a follicle measuring 4.0 mm. The outcome of insemination could not be determined in 5/6 animals that had not yet ovulated at the time of assessment. A large blood clot was found in the uterus on the side of insemination in two animals, one of which had the sperm-covered egg in the lower oviduct.

Sperm and Egg Transport After IUAI

Spermatozoa were deposited into the ipsilateral uterus but were recovered from all sections of the female tract within 0.4 h and, with the exception of the upper oviduct, remained there for up to 6.2 h (Table 2). No spermatozoa were recovered between the uterus and upper oviducts on the contralateral side. The highest numbers of spermatozoa were located in the anterior lateral vaginae at 0.4 h and in the uterus 6.2 h after insemination. Recovery of spermatozoa by flushing decreased from 8.7% to 1.6% 6 h later. Tissue from the median vagina to the urogenital sinus was not assessed for the 1.2-h animal and thus percentage recovery could not be calculated. Within 6.4 h after IUAI, one female had ovulated, spermatozoa had reached the oocyte, and the egg had migrated down to the lower oviduct.


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  		TABLE 2. Number of spermatozoa recovered from the female reproductive tract 0.4–6.2 h after IUAI in the tammar wallaby

Progesterone and Estradiol Concentrations

There was no relationship between either progesterone or estradiol levels and the time of insemination after birth (r = 0.075, P = 0.637, n = 42 and r = –0.218; P = 0.155, n = 44, respectively) (Fig. 4). In the two females that gave birth (early and late IUAI), progesterone values at the time of insemination were 331.2 and 214.3 pg/ml and estradiol values were 38.6 and 39.1 pg/ml, respectively. In the female that had ovulated a sperm-covered oocyte, progesterone levels dropped from 507.9 pg/ml at insemination to 357.1 pg/ml at autopsy 6.4 h later. Estradiol levels in this female changed little, from 30.5 pg/ml to 34.3 pg/ ml, over the same period.



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  		FIG. 4. Steroid hormone levels of (a) progesterone and (b) estradiol-17ß at the time of AI in 46 female tammar wallabies. Black rectangles, birth of pouch young; black circle, sperm-covered oocyte; white circle, unsuccessful; white triangles, yet to ovulate

Microsatellite Paternity Analysis

Allele sizes of each offspring were consistent with the inheritance of one allele from the known mother and a second allele from the respective AI sperm donor (Table 3). After determining the frequency of alleles in our captive colony, Cervus 2.0 software identified the first AI sperm donor as the most likely candidate father of one AI offspring with 95% confidence (Log of the odds ratio (LOD) = 4.3, {Delta} = 2.6). Based on the known maternal genotype, the probability of a false-paternity assignment was 0.04%. The second AI sperm donor was identified with 80% confidence as the most likely candidate father for the second AI offspring (LOD = 4.2, {Delta} = 1.7). Again, based on a known maternal genotype, the probability of a false paternity assignment was 0.19%.


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  		TABLE 3. Allele sizes of known mother, pouch young (PY), and the corresponding AI sperm donors and paternity assignment scores for the two tammar wallaby offspring born following artificial insemination


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
This study demonstrates that IUAI can produce live macropodid marsupial young. Of the 22 animals inseminated by IUAI in which an outcome could be determined, two gave birth and, in one, sperm had reached the egg. No births resulted from any of the other sites of insemination. This success rate is comparable with 4–10% IUAI birth rates per cycle in humans [27, 28], but well below 40–70% AI pregnancy rates regularly achieved in dairy cattle [29, 30]. The relatively high success rate reported in the koala may be due to a prolonged estrus and the synchronization achieved as a result of ovulation induced by seminal factors [31]. In most other marsupials, however, ovulation is spontaneous and fertilization occurs long after mating [7].

After superovulation, fertilization success in the possum and tammar wallaby using AI is high [3]. However, more than 60% of embryos recovered from superovulated possums 4 days after IUAI had degenerated [2]. Mucin and shell coats isolated from oocytes and early embryos of these superovulated possums were thinner and had an altered protein composition [13]. These egg coats appear critical during preimplantation embryo development and blastocyst expansion [11]. From the results of recent attempts at AI [2–5, 32], it appears that superovulation may affect embryo viability in monovular marsupials, possibly by altering the structural integrity of the tertiary egg coats.

Our study has shown that both electroejaculated and epididymal spermatozoa deposited into the uterus are capable of oocyte penetration and fertilization and of supporting normal embryo development to term. Fertilization after IUAI using either electroejaculated or epididymal spermatozoa have been reported previously in the possum and tammar [2]. In addition, possum spermatozoa recovered from the oviducts at least 4.5 h after IUAI or cocultured with oviduct epithelial cell monolayers were capable of zona binding and penetration in vitro [33] Thus, in the tammar, capacitation appears to occur in the oviduct, as it does in eutherian mammals [34].

After IUAI, we found spermatozoa in the upper oviduct within half an hour but not after 6 h (n = 4). In the possum, sperm numbers in the ampulla of the oviduct were highest at 6 h and declined significantly 12 h after IUAI [32]. Possum spermatozoa may require at least 4.5 h in the oviduct to be capable of zona penetration [33] and, in superovulated tammars, maximum fertilization rates are achieved after IUAI 4–7 h before expected ovulation [2, 3]. Thus, in the tammar, it appears that spermatozoa should be delivered by IUAI 4–6 h before ovulation to maximize the number of spermatozoa at the site of fertilization and allow sufficient time for capacitation to occur. In this study, one successful female had ovulated, spermatozoa had reached the oocyte in the upper oviduct before mucin deposition, and the egg had migrated to the lower half of the oviduct in less than 6.4 h. Such rapid oviductal transport is a feature of marsupial reproduction [7, 10]. Timing of AI is critical in marsupials both because fertilization must occur before the mucin and shell coats are laid down and passage of the oocyte through the oviduct is very rapid [8].

In our study, inseminations into different sites of the female tract were conducted at times similar to those in which spermatozoa are found in these regions after natural mating [16], but most were unsuccessful. During natural mating, the female typically mates with several males and the total volume of semen in the tract often exceeds 80 ml [16]. The coagulated ejaculate may act as a reservoir from which spermatozoa migrate and replenish spermatozoa in the upper tract [16]. In contrast, AI deposits spermatozoa into a specific region of the tract without further replenishment. Thus, timing of AI may need to be closer to ovulation than for natural mating so that spermatozoa do not dissipate or break down before ovulation.

Breakdown and loss of sperm may contribute to low insemination success. After IUAI, up to 30% of spermatozoa are excreted retrogradely in the pig [35] and are detected in the vaginal smears of tammars and possums [2]. Tammar spermatozoa are also broken down in the tract by phagocytosis [16]. In our study, the sperm-rich medium delivered by catheter to the anterior vaginal culs de sac may have been lost back through the median vagina because the volume (2 ml) and viscosity were low compared with the large coagulating ejaculates seen after natural mating [16]. In addition, we found spermatozoa migrated both anteriorly and posteriorly shortly after IUAI and sperm recovery fell to less than 2% after 6 h in the tract. However, in our study, the total number of spermatozoa inseminated into the uterus and later recovered in the lower oviduct was equal to or up to 100-fold more than that found in these regions after a single natural mating [16]. Further, the three successful females were inseminated with low numbers of spermatozoa relative to unsuccessful animals and so, at least in the IUAI-treated females, it is unlikely that breakdown or loss of spermatozoa led to reduced success.

Of the three AI attempts considered successful, the two females that were inseminated late received spermatozoa of poor motility while the female inseminated early received very high-motility spermatozoa. These two animals were inseminated closer to the predicted time of ovulation than most other animals treated. Thus, IUAI with spermatozoa of poor motility can still result in births if the timing is close to ovulation, but the use of spermatozoa with high motility may allow for an earlier time of insemination. However, insemination of sperm with strong motility is critical in the lower tract, as spermatozoa are required to navigate more barriers and travel further to the site of fertilization [34]. Thus, the use of spermatozoa with poor progressive motility for the urogenital sinus insemination may partially explain the failure of this group in our study.

In our study, at the time of examination, one female (49.4 h p.p.) had ovulated and six animals (43–60 h p.p.) had not ovulated during the ovulation window determined previously [10]. The largest Graafian follicle observed was 4.9 mm, which was equal to the maximum seen in this species (M.B.R., unpublished data). We found no relationship between circulating concentrations of plasma progesterone or estradiol and time after birth, and the success of fertilization was not associated with any pattern in hormone levels.

Surgical procedures may compromise AI success. Anesthesia during proestrus prevents an increase in plasma LH and delays ovulation in sheep [36], blocks ovulation in the domestic cat [37], and compromises sperm transport in vaginally inseminated felids, canids, and mustelids [reviewed in 38] but not the brushtail possum [5, 32]. In the tammar, different combinations of anesthetic and laparotomy have a detrimental effect on survival of fetuses in early but not late pregnancy [39]. Similarly, laparoscopy is thought to affect ovulation and egg transport to the oviduct in the possum [2, 40]. In our study, compromised sperm transport due to anesthesia may explain the failure of the 14 animals inseminated into the urogenital sinus and anterior vaginal culs de sac. However, we observed rapid sperm and egg transport through the oviduct after IUAI using anesthesia and laparotomy, suggesting that, at least for intrauterine inseminations, these procedures may have had little effect on ovulation, transport, and fertilization.

In our study, more extensive surgery was required to expose the cervices for transcervical inseminations, which may have compromised ovulation or pregnancy in these animals. In addition, uterine blood clots found on the side of the IUAI in two females at autopsy (one containing the sperm-covered oocyte) suggest this may have also occurred in some animals left to give birth. While gamete transport and fertilization appear unaffected, it is not known if the clots may create a uterine environment detrimental to embryo survival.

Excessive handling of animals can affect the timing of estrus and, in tammars regularly handled on a 4- to 8-h basis, mating fails to occur or is delayed to 10 h p.p. [16, 23], compared with 1.3 h p.p. in free-ranging animals observed without disturbance [10, 14]. In our study, animals were only checked every 12 h, but this in combination with insemination procedures may have been sufficient to delay estrus and possibly ovulation.

In summary, because of the low success rate, the relationship between sperm quality, insemination route, and timing of insemination remains unclear. However, this study has demonstrated the potential for IUAI to generate viable offspring using epididymal or electroejaculated spermatozoa in the tammar wallaby. Further studies are now needed to optimize the procedures.


   ACKNOWLEDGMENTS
 
We thank Dr. David Parer for the photograph of the tammar neonate (Fig. 1a) originally published by Surrey Beatty and Sons Pty. Ltd. [19]. We thank all members of the wallaby research group and, in particular, John Akamatis, Scott Brownlees, Kerry Martin, and Sue Osborn for assistance with animals. Jan Loose kindly performed the estradiol and progesterone assays. Insemination catheters, media, and bromocriptine were kindly donated by Cook Australia, Invitrogen, and Novartis, respectively. Microsatellite analysis was kindly subsidized by the Commonwealth funded AGRF, and Cervus 2.0 paternity assignment software was provided by Tristan Marshall.


   FOOTNOTES
 
1 Supported by a SPIRT grant (C09930004) from the Australian Research Council to M.B.R., G.S., P.T.-S. and Zoos Victoria, and a Ph.D. scholarship from the University of Melbourne Faculty of Science to D.B.B.P.P. Back

2 Correspondence. FAX: 61 3 93481719; m.renfree{at}zoology.unimelb.edu.au Back

Received: 28 June 2004.

First decision: 12 July 2004.

Accepted: 2 September 2004.


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 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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