New Reproductive Technologies For The AI Industry

Donald G. Levis, Ph.D.
Ohio Pork Industry Center, The Ohio State University, Columbus, OH 43210


The techniques and application of artificial insemination (AI) have greatly improved since it was first suggested as a viable technique to inseminate sows in 1948 (Ito et al., 1948). Although scientists promoted the use of AI of pigs in 1956 and 1966 (Polge, 1956; Melrose, 1966a,b), the worldwide pork industry did not rapidly adopt the use of AI. The United States pork industry has dramatically increased the use of AI during the last few years. The rapid increase in adoption of AI is most likely a result of a substantial improvement of AI techniques, AI equipment, genetic evaluation programs, measurement of carcass merit techniques and payment for carcass merit by packers (Singleton, 2001). A survey of swine operations within the 17 leading pork-producing states (representing nearly 94% of swine herds with 100 or more pigs on December 1, 1999) found that 85.3% of sows were artificially inseminated when the enterprise had more than 500 breeding females (USDA, 2001). The reproductive technologies briefly discussed in this paper are boar semen extenders, intrauterine insemination of female pigs, sexing boar semen, and cloning pigs.

Boar Semen Extenders

The chemical composition of boar semen extenders developed prior to 1991 was recently presented in a review by Levis (2000). Today, the chemical composition of “new and improved” long-term liquid boar semen extenders is not revealed because of the large amount of worldwide competition among companies manufacturing and marketing boar semen extenders. As a result of the rapid adoption of AI, a huge market exists for liquid boar semen extenders and a need for long-term (7 to 10 days) extenders. Some of the reasons for developing new long-term boar semen extenders relate to: (1) more favorable work hours in the boar stud, (2) reduced transportation cost of delivering semen to the farm, (3) testing of semen for Porcine Reproductive and Respiratory Syndrome virus before use, and (4) a longer period of time for using the semen with satisfactory reproductive performance

University scientists in the United States are not actively involved in the development and field evaluation of new long-term semen extenders. Currently, the data generated on long-term viability of sperm cells, farrowing rate and litter size are primarily by the company with a vested interest in the new long-term (10 to 15 days of storage) boar semen extender. The most recent data on “new” extenders developed by companies marketing semen extenders is not peer reviewed or published (most likely will never be published) in scientific journals. However, this information is utilized in an aggressive advertisement and marketing campaign. Very few boar studs or pork production enterprises compare in a scientifically manner “new” extenders to existing extenders. It is important for the well being of the pork industry that companies who market boar semen extenders generate scientifically sound data on evaluation of extenders before the extenders are marketed. Several pork production enterprises believe they have adequately compared the reproductive performance between two extenders; however, many times they have confounded type of extender with ejaculate and week of semen collection. To clarify, the pork production unit use Boar A and Extender A during Week 1 and Boar B and Extender B during Week 2 or use Extender A during Week 1 and Extender B during Week 2. These type of comparisons can often lead to false interpretation of results.

The most recent “advertisement” data on the new long-term boar semen extenders can be found at the company locations listed below. The experimental protocol is not presented for any of the field data.

Minitube web site -
Title of article - EnduraGuard? Performance Proven in Field Trials
IMV web site –
Title of article – SafeCell Plus? An Innovative Concept in Long Term Swine Semen Extenders

Intra-uterine Insemination

The current protocol for inseminating pigs is to: (1) check for estrus once or twice per day, (2) inseminate females two to three times during estrus, (3) inseminate 2.5 to 5.0 billion sperm cells per dose, (4) use a total volume of 80 to 100 mL, and (5) deposit the semen into the caudal to middle segment of the cervix with a disposable catheter. The site of semen deposition, number of sperm cells per dose, volume per dose of semen inseminated, time of ovulation and number of inseminations per estrus are the main factors that influence the current protocol for inseminating pigs with liquid-extended semen. To increase the efficient use of spermatozoa from a single ejaculate of boar semen, “new protocols” for using liquid-extended semen will utilize a smaller volume, a reduction in number of spermatozoa per dose and change the site of semen deposition. One method to enhance the number of female pigs that can be inseminated by each ejaculate of boar semen is to use intrauterine insemination. There are basically three procedures for depositing spermatozoa into the uterus: First, surgically deposit spermatozoa approximately 5 cm (?2 inches) from the uterotubal junction (Krueger et al., 1999; Krueger, 2000; Krueger and Rath, 2000; Rath et al., 2000); Second, nonsurgically deposit semen into the uterine body (Anonymous, 2001; Levis et al., 2002; Watson, 2001); and Third, nonsurgically deposit semen “deep” into the uterine horn (Martinez et al, 2001, 2002).

Intrauterine Body Insemination (IUBI)

The uterine body of the pig, approximately two inches long, is located between the cervix and bifurcation of the two uterine horns. When using IUBI, semen is deposited about 20 cm (8”) farther into the female pig’s reproductive tract compared to traditional cervical AI (Figure 1). The use of IUBI does not overcome the “biological” factor of losing sperm cells during the transportation process from the uterine body (site of semen deposition) to the oviduct (site of fertilization). Sperm cells are primarily lost by back-flow of semen during the first two hours after AI and phagocytosis by polymorphonuclear (PMN) leucocytes. Although approximately 90% of the spermatozoa cannot be recovered from the uterus within 2 hours after a natural mating (Viring, 1980), a sufficient number of sperm cells (100 to 200 million) reach the uterotubal junction and the adjacent first isthmic segment of the oviduct (sperm reservoir) within 15 to 20 minutes after a natural mating (Hunter, 1990). Because a substantially lesser number of motile sperm cells are deposited in the female reproductive tract with AI (2.5 to 3.0 billion) compared with a natural mating (47.9 ± 13.6 billion; ejaculation interval of 3 to 4 days), it is extremely important to minimize the number of spermatozoa lost during the transport of spermatozoa from the site of semen deposition to site of fertilization after AI.


Steverink et al. (1998) documented that back-flow is frequently observed during the period when cervical insemination is being performed (63.3% of sows) and 0 to 30 minutes after (98.2% of sows) cervical insemination; plus, volume of back-flow is quite variable during insemination (? = 7%, range is 1 to 56%) and 0 to 30 minutes after insemination (? = 31%, range is 3 to 76%). Although the causes of back-flow among sows are still poorly understood, many times the amount of back-flow is influenced by skill level and patience of the inseminator (technician).
An important question is: Does back-flow affect farrowing rate and litter size born alive? For back-flow to have a significant affect on farrowing rate and litter size, there has to be a loss of sperm cells as a consequence of back-flow. Steverink et al. (1998) found a high linear correlation between volume of “fluid” lost and number of spermatozoa lost (80 mL dose with 1, 3 or 6 billion sperm) during insemination (r = .97), 0 to 30 minutes after insemination (r = .73) and 30 minutes to 2.5 hours after insemination (r = .81). The percentage of total spermatozoa lost during insemination was 8% (range, .3 to 50%) and 14% (range, .3 to 79%) at 0 to 30 minutes after insemination. Matthijs et al. (2000) found that 45% of the spermatozoa inseminated (80 mL dose) were recovered in back-flow fluids collected in a stoma bag attached around the vulva.

The experiment by Steverink et al. (1998) did not allow the sows to farrow; thus, farrowing rate could not be calculated. The experiment did evaluate the effect of backflow on fertilization rate of oocytes. Although the number of observations for the large volume of fluid backflow category was very small (n = 4 to 5 sows), a negative effect was found when a large volume of back-flow occurred during the insemination process with one billion sperm on the percentage normal embryos. The percentage of normal embryos was reduced (P < .05) regardless of the interval from time of insemination to ovulation. There were not an adequate number of observations to evaluate whether the interval of time from insemination to ovulation has an effect on percentage of normal embryos. Numerically, 68% of the embryos were normal for sows inseminated with 1 billion sperm cells at 0 to 24 hours before ovulation compared with 46% normal embryos of sows inseminated with 1 billion cells at 24 to 48 hours before ovulation. The amount of back-flow after insemination did not affect the percentage of normal embryos with any of the three insemination dosages (1, 3 or 6 billion).


Woelders and Matthijs (2001) have reviewed the scientific literature on phagocytosis of boar spermatozoa in vitro and in vivo. Uterine clearance of “foreign” material is a normal physiological process that serves to prepare the uterus for the reception of embryos. The clearance of spermatozoa (phagocytosis) from the reproductive tract by PMNs is not a specific immune response; otherwise, insemination would lead to the development of a sterilizing anti-sperm immunity reaction.

Insemination of pigs triggers a massive influx of PMNs into the lumen of the uterus. Large numbers of PMNs have been found at 30 minutes (Lovell and Getty, 1968), 2 hours (Pursel et al., 1978), and 3 hours (Kohsaka et al., 2000) after insemination. Rozeboom et al. (1999) found greater numbers of PMNs from 12 to 36 hours after sows were inseminated with 5 billion sperm cells compared to sows inseminated with 100 mL of seminal plasma or phosphate buffered saline. Because phagocytosis in the uterus kills sperm cells, it is extremely important that sperm cells travel to the oviduct as quickly as possible. Although the sperm cells do not travel through the folds of the cervix with IUBI, they do have to travel through the long, convoluted structure of the uterine horns.


The reproductive performance in various studies that utilized IUBI is indicated in Table 1 and 2. Most of the studies have confounded results due to the differing types of inseminating catheter, volume of semen and number of sperm cells per dose; thus, the “main” effect of type of catheter on reproductive performance is unknown. In most of the studies the IUBI catheter has been mainly used as a “tool” to by-pass the folds of the cervix; thus, it is anticipated that a lesser volume and fewer sperm cells per dose are needed. Although values for farrowing rate and litter size born live are routinely reported, a fecundity index value (farrowing rate x litter size) provides a more reliable estimate for determining the value of IUBI procedure. A decision for adopting a new reproductive technology should not be based on a single trait, such as, farrowing rate or litter size. When making a comparison between “treatments”, one frequently finds that Treatment 1 resulted in a greater farrowing rate compared with Treatment 2 but utilization of Treatment 1 resulted in a lesser litter size than Treatment 2.

Computer Model. In order to evaluate the economics surrounding the use of IUBI, a Microsoft Excel spreadsheet was developed. The spreadsheet can simultaneously evaluate three scenarios. This spreadsheet is available from the Ohio Pork Industry Center’s website ( The farrowing rate and litter size born live data from Gil et al. (2002) was used to generate the economics of using intrauterine body insemination (Table 3). The basic factors used in the model are farrowing rate, litter size born live, weekly farrowing, number of farrowing crates per group per week, percentage preweaning death loss, duration of time (minutes) per insemination, dollars per hour of labor, percentage of group is gilts, cost per each type of insemination catheter, cost per dose of semen, number of inseminations per estrus, and assumed profit per pig. This model is designed to have a specific number of sows bred based on the estimated farrowing rate (over-breed); thus, all assigned farrowing crates per group are filled to capacity.

When a dose of semen is the same price (regardless of number of sperm cells per dose) for the data by Gil et al. (2002), the model indicated an economic loss (-$615 to -$79,693) in eight of the ten trials when IUBI was used compared with cervical AI (Table 3). If the price per dose of semen is $3.00 for IUBI and $6.00 for cervical AI, seven of the ten trials showed an economic advantage that ranged from $3,066 to $51,078 for IUBI. Using the data generated by Watson and Behan (2001), the cost of IUBI semen needed to be about $1.00 less per dose to produce the same yearly net profit as cervical insemination (Table 4). The results from these trials clearly demonstrate that the price per dose of semen plays a critical role in the economical use of IUBI. What will semen suppliers charge per dose of semen that contains one billion or fewer spermatozoa? It must be remembered that a dose of semen is priced as a combination of genetic cost, number of sperm cells per dose, overhead costs, and profit. These factors will determine the differential price of semen doses that contain greater or lesser sperm numbers.

Deep Intrauterine Horn Insemination (DIUHI)

To further reduce the number of spermatozoa per dose inseminated, techniques are being developed whereby sperm cells are place deep into the uterine horn at a lesser dosage volume as compared with that used with IUBI (Martinez et al., 2001; Rath, 2002). Although it is not practical to surgically deposit sperm cells into the uterine horn on a commercial farm with the presently available technologies, it has been documented that fertilization potential is not substantially decreased when 100 million sperm are deposited about 5 cm (? 2 inches) distal from the uterotubal junction (Krueger et al., 1999; Krueger and Rath, 2000; Rath et al., 2000). A field experiment in the United States by Krueger and Rath (2000) did find a non-significant linear decrease in number of piglets born live as number of spermatozoa per dose decreased (Table 5). The difference detected might be significant when a larger number of sows per treatment are used. The encouraging results from depositing much reduced spermatozoa close to the uterotubal junction on farrowing rate and litter size has stimulated scientists to investigate non-surgical, non-sedative methods of depositing spermatozoa deep into the uterine horn.

Fiber Optic Insemination

The anatomy of the cervix (thick muscles, series of folds or ridges, cervical contractions during estrus) and long uterine horns (120 to 140 cm [47 to 55 inches] long with convoluted structures) previously impeded the development of a procedure for non-surgical insemination into the uterine horn. A fiber optic endoscope technique for non-surgical DIUHI of pigs has been investigated in Spain (Martinez et al., 2001). Table 6 provides data that indicates the effect of number of sperm cells per dose on reproductive performance when inseminating non-sedated sows with a flexible fiber optic endoscope. Farrowing rate and number of piglets per litter were not significantly different when sows were inseminated using fiber optic technologies with 1 billion, 200 million or 50 million sperm cells as compared to cervical AI with 3 billion cells. Numerically, the number of piglets per litter was fewer on the fiber optic treatment. The lack of detecting a significant difference might be due to the small number of sow (13 to 18 sows) per fiber optic treatment group.

Flexible Catheter Insemination

Although estrus sows can be successfully inseminated with a fiber optic endoscope, the endoscope is expensive, fragile, and most likely unsuitable for use under field conditions. Scientists in Spain have evaluated a specifically designed flexible catheter (1.8 m [70 inches] long) that is inserted through a traditional spirette catheter and passed through the cervix and moved forward along ONE uterine horn until its total length has been inserted to about the middle of the uterine horn (Martinez et al., 2002). In this study, crossbred sows were treated with 1250 IU equine chorionic gonadotrophin (eCG) 24 hours after weaning and with 750 IU of human chorionic gonadotrophin (hCG) 72 hours after eCG. DIUHI was performed once at 36 hours after hCG treatment with 150 million, 50 million, 25 million or 10 million sperm cells in 10 mL. Control sows were cervically inseminated twice with 3 billion sperm cells in 100 mL. Farrowing rate after DIUHI with 150 million or 50 million sperm cells did not differ from the control group (Table 7). However, farrowing rate was less (P < .001) after DIUHI with 25 million or 10 million sperm cells compared with control sows. Although litter size born was not significantly different between treatments, litter size was numerically smaller for sows inseminated with 10 million, 25 million or 50 million sperm cells.

Ipsilateral Fertilization

Because sperm cells are only deposited in one uterine horn, the question arises as to whether fertilization only takes place in the uterine horn where spermatozoa are deposited (ipsilateral fertilization) or whether fertilization also takes place in the opposite uterine horn of spermatozoa deposition (contralateral fertilization). Research has demonstrated that when spermatozoa are deposited close to the uterotubal junction in one uterine horn, spermatozoa are able to reach the contralateral oviduct and fertilize the oocytes (Martinez et al, 2002). The total number of normal embryos was fewer in the assumed “contralateral” uterine horn (Table 8). The mechanism by which sperm cells are transported to the contralateral oviduct is being studied. Hunter (1978) reported that fertilization of oocytes occurred after intraperitoneal deposition. In addition, Viring and Einarsson (1981) suggested that spermatozoa pass through the oviduct of pigs into the abdominal cavity during the first hours after natural mating. Yaniz et al. (2002) recently reviewed the scientific literature on intraperitoneal insemination in mammals.

Uterine Infection

If sows are inseminated during estrus, it has been suggested that uterine infections (vaginal discharges) is less than 1% (Martinez et al, 2002). The low risk of inducing uterine infection with DIUHI is most likely because sows are resistant to bacterial infections when circulating concentration of estrogen is elevated during estrus (De Winter et al., 1996). Vaginal discharges will be a problem if good estrous detection procedures are not utilized to prevent the insemination of anestrous sows or sows close to going out of estrus.

Animal Welfare

Will animal welfare activist accept the DIUHI procedure? Is the DIUHI procedure painful to the sow? Martinez et al. (2002) studied the difficulties encountered during insertion of the flexible catheter, duration of time to insert catheter and behavior of the sow during insertion. The flexible catheter was successfully placed into one uterine horn in 95.4% of the sows in an average of 3.7 ± .09 minutes. Parity (2, 3 or 4 to 6) did not significantly affect the difficulties or time required to insert the flexible catheter. The percentage of sows expressing good or moderate behavior during the insemination procedure was 97.1% when there was no or minor difficulty at insertion of catheter, 93.8% when there was medium difficulty at insertion of catheter, 85.7% when there was high difficulty at insertion of catheter and 94.4% when it was impossible to insert catheter.


At this point in time it is impossible to economically evaluate the use of DIUHI. The cost of semen and insemination device is unknown in the United States. MAGAPOR has a DIUHI device (FirFlex?) on the market in Europe ( Fewer boars are needed to produce semen for cervical artificial insemination compared with natural service. Fewer boars are needed to produce semen for DIUHI compared with cervical insemination. Fewer boars are needed to produce semen for in vitro fertilization of ova (75 sperm cells per oocyte; Rath et al., 1999) compared with DIUHI. How many boars will be needed to produce sperm cells for use with DIUHI? Will the genetic companies have the correct boars identified to produce terminal and maternal semen?

Number of Boars Producing Semen. Scientists have made statements that: (1) DIUHI will be of great importance to the pork industry because superior boars can be more efficiently utilized, (2) DIUHI will complement the development of AI, especially with the use of sexed semen, (3) DIUHI will allow a tremendous saving in cost of semen. However, the use of a lesser number of sperm cells per insemination will have a significant influence on genetic companies.

The influence of number of sperm cells per dose and genetic line on total number of doses packaged is indicated in Table 9. An estimated number of “productive” boars required in the United States for sperm production when servicing sows by natural service, cervical artificial insemination or DIUHI is indicated in Table 10. The number of boars needed to produce spermatozoa for use with DIUHI is about 890 (150 million sperm per dose) to 5,900 (1 billion sperm per dose). If only 890 boars are required, who will own this small number of boars?

Sexing Boar Semen

The implementation of specific mating systems to produce gilts (maternal line) or boars (terminal line) and separate-sex feeding programs has increased the need for sexed semen. If pork producers are marketing gilts, the male pigs produced are “by-products”. Conversely, if pork producers want a population of male pigs, the female pigs produced are “by-products”. The only method of achieving the desired sex of offspring is to separate the X-chromosome from the Y-chromosome bearing spermatozoa before fertilization of ova. Attempts to separate X and Y spermatozoa have been based on sperm head volume (van Munster, 2002; van Muster et al., 1999), swimming velocity (Beernik et al., 1993; Penfold et al., 1998), the Y-linked histocompatibility antigen (H-Y antigen) expressed on the surface of male somatic cells (Hendriksen et al., 1993), plasma membrane proteins (Blecher et al., 1999; Hendriksen et al., 1996; Hendriksen, 1999), time of insemination (Rorie, 1999), and DNA content (Johnson, 2000; Rens, W. et al. 1999). The method that has been repeatedly proven to be effective in producing offspring of the predicted sex is the Beltsville Sperm Sexing Technology (BSST). This method separates sperm cells on the basis of DNA content using flow-cytometric sperm sorting.

Brief Description of Beltsville Sperm Sexing Technology

Scientists have described the BSST procedure and validation of desired offspring (Guthrie, et al., 2002; Johnson et al., 1999; Johnson and Welch, 1999; Welch and Johnson, 1999). The current BSST uses a flow cytometer/cell sorter (MoFlo) modified specifically for spermatozoa to measure the relative florescent intensity reflective of DNA content of EACH spermatozoon as it traverses a laser beam (125 mW; 1 to 2 ?second) and then separates the sperm based on DNA content. The Y chromosome of boar sperm is smaller and carries 3.6% less DNA than the larger X chromosome. The dye used to stain the DNA of living sperm is a bisbenzimide fluorescent vital dye (Hoechst 33342) sensitive to ultraviolet light. Proportion of stain per number of spermatozoa is critical to maintain uniform staining (8 ?L Hoechst 33342 [5 mg/mL stock] per one mL aliquot of semen with 150 million sperm cells). The suspension is then incubated at 35?C for approximately one hour. Currently, 2 mL of sperm are prepared with the appropriate amount of stain and BTS extender to give 200 to 300 million sperm for sorting over about a 2-hour period. Just prior to sorting, the stained sperm are treated with FD&C#40 food coloring for five minutes. The food coloring penetrates the membrane of the dead sperm and quenches the intensity of fluorescence of the dead sperm, thus eliminating dead sperm from the viable sperm population.

The high speed MoFlo cell sorter, modified for sperm sorting and includes an orienting nozzle, can produce six million X sperm (96 to 98% purity) and six million Y sperm (93 to 95% purity) per hour. When only sorting for X sperm, the sorter can produce 11 million sperm per hour (85 to 90% purity). The high-speed sperm sorting system with orienting nozzle is capable of processing more than 40,000 events per second (144 million events per hour). Orientation of sperm cells at time of passing through the laser beam is one aspect that limits the output of “purified” populations of X and Y spermatozoa. The current beveled needle with orienting nozzle correctly orients (edge of sperm cell oriented to the 90? fluorescence detector and flat side oriented to 0?fluorescence detector) about 70% of the sperm cells.

Farrowing Rate and Litter Size

Research on fertility of sexed semen has been in combination with: (1) in vitro maturation and in vitro fertilization of ova followed by surgical transfer of embryos into the oviduct [IVM-IVF-ET], or (2) surgical placement of spermatozoa in the oviduct. The results of these experiments are indicated in Table 11. Although the BSST is capable of producing sexed sperm for use with IVM-IVF-ET or surgical insemination to produce offspring of the desired sex, farrowing rate and litter size is not acceptable for commercial use.

Implementation of Technology

Although X and Y boar spermatozoa can be successfully separated on DNA content, I believe the commercial application of sexed semen on a large number of sows is several years away. As with any new reproductive technology, the use of sexed semen will depend on efficiency, economics, effectiveness, and ease of use. For commercial introduction of flow-sorted sexed semen, new instrumentation and procedures must provide: (1) a greater sort rate, (2) adequate viability of spermatozoa during transportation, (3) long-term storage that is equal to the current storage time of liquid extended semen, (4) a method to inseminate females with a smaller volume and lesser number of sperm cells [such as, DIUHI], and (5) a product that produces an adequate farrowing rate and litter size. Undoubtedly, the cost of sexed semen will be greater as compared with non-sorted semen; thus, only high-value sexed sperm cells should be marketed.

Cloning and Embryo Transfer of Pigs

Major reasons why scientists are interested in cloning pigs include: (1) future production of pig organs that can be successfully transplanted into humans (Dai et al., 2002; Lai et al., 2002; Polejaeva, 2001), (2) to study human diseases (Petters, 1994), and (3) to increase dissemination of superior genetics. Although the production and embryo transfer of cloned piglets is now a reality (Betthauser et al., 2000; Bondioli et al., 2001; Dai et al., 2002; Lai et al., 2002; Onishi et al., 2000; Polejaeva et al., 2000), the reproductive performance of surrogate sows is extremely inadequate (Table 12). Because of the complexity and greater cost of biotechnology procedures used to produce cloned pigs (Figure 2), it will be several years before “commercial companies” will be providing cloned pigs on a large scale to the pork industry. Currently, some biotechnology companies involved with cloning livestock are having difficulty in obtaining funding to develop these technologies (Nature 418:903, August 2002). Biotechnology companies have little interest in cloning pigs for genetic merit of production traits. The main objective of biotechnology companies is to produce pigs that can be utilized to enhance human health aspects. Dr. Michael Bishop, former head of Infigen, recently stated: “I do not think that livestock cloning will ever be profitable”.

If cloning of pigs becomes financially feasible, where would the technology benefit the area of artificial insemination? The most logical aspect is sperm production and boar fertility. It is well known that genetic progress is more rapidly accomplished by using genetically superior boars compared to sows. It is well documented that boars differ in their ability to produce spermatozoa and fertilize ova (Flowers 1998-2000, 2001, 2002; Xu et al., 1996a,b; Xu et al., 1998). During 2001, Infigen cloned two boars that produced show-pig semen for Prairie State Semen, Inc. ( Research needs to be conducted to evaluate whether enhanced sperm production and fertility are maintained in boars cloned from superior sires.

Table 1. Reproductive performance of sows artificially inseminated by intrauterine body insemination (IUBI) or traditional insemination (cervical)

a Gil et al., 2002 (volume of IUBI is = 50 mL; volume of cervical is 90 mL)
b Levis, et al., 2002 (volume of IUBI with 1 billion is 30 mL; volume of IUBI with 1.5 billion is 50 mL; volume of cervical with 3 billion is 100 mL)











Figure 1. Placement of intrauterine catheter into uterine body of a pig

Figure 2. Production of clones with a double nuclear transfer procedure (Nature 407:86-90)



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