Laparoscopic Ovum Pick-up For In Vivo Oocyte Retrieval In Alpacas

Summary

This protocol describes a reproducible method for in vivo oocyte recovery in alpacas using hormonal synchronization, FSH superstimulation, and laparoscopic ovum pick-up (LOPU). The procedure yields consistent follicular responses and efficient oocyte recovery, supporting applications for in vitro embryo production and advanced reproductive biotechnologies.

Abstract

Alpacas are increasingly incorporated into reproductive and biomedical research due to distinctive physiological and immunological characteristics that extend their relevance beyond agricultural applications. As camelids, alpacas exhibit induced ovulation, prolonged follicular dynamics, asymmetric uterine physiology, and a unique immune system characterized by heavy-chain–only antibodies. These features support their use in comparative and translational research, particularly in reproductive biology and genetic engineering. Laparoscopic ovum pick-up (LOPU) is a minimally invasive approach for oocyte recovery in species with complex reproductive anatomy, allowing direct ovarian visualization, precise follicular aspiration, and reduced tissue trauma compared with transvaginal techniques. The objective of this study was to develop and standardize a reproducible in vivo oocyte retrieval protocol in alpacas using LOPU. Eight adult female alpacas (3–8 years of age) were enrolled in this study and subjected to repeated LOPU sessions conducted once per month throughout the study period. Alpacas were evaluated by transrectal ultrasonography, and animals presenting ovarian follicles ≥7 mm received 50 µg gonadotropin-releasing hormone intramuscularly. Ovarian superstimulation was induced using a total dose of 200 mg follicle-stimulating hormone administered in a decreasing twice-daily regimen over four days. LOPU was performed 12–14 h after the final hormone administration under general anesthesia induced with ketamine, xylazine, and butorphanol and maintained with isoflurane. A three-port laparoscopic approach with carbon dioxide insufflation was used. Follicles were aspirated using a 20-gauge needle connected to a regulated vacuum system (25–30 mmHg; 13–15 mL/min). Oocytes were recovered in TCM-199 medium and evaluated under a stereomicroscope. Alpacas exhibited a mean of 14.88 ± 3.85 follicles, with 10.94 ± 3.62 follicles aspirated per session and 6.68 ± 2.21 oocytes recovered, corresponding to a recovery rate of 61.30 ± 8.21%. These findings suggest that LOPU may represent a viable approach for in vivo oocyte retrieval in alpacas, with potential applications in reproductive biotechnology workflows.

Introduction

South American camelids, particularly alpacas (Vicugna pacos), have gained increasing relevance in reproductive biotechnology due to their distinctive reproductive physiology and expanding role in biomedical and agricultural research. Alpacas exhibit induced ovulation, prolonged follicular waves, and unique ovarian dynamics that differ markedly from those of domestic ruminants¹. These characteristics, together with the growing economic and scientific importance of the species, have driven efforts to develop assisted reproductive technologies (ARTs) tailored to camelid biology. Nevertheless, progress in alpaca ARTs has historically lagged behind that achieved in cattle, sheep, and goats, largely owing to anatomical constraints, limited access to high-quality gametes, and technical challenges associated with in vivo oocyte recovery²˒³.

Early attempts to recover oocytes from live alpacas relied primarily on surgical laparotomy, a method adapted from foundational studies in South American camelid embryo research³˒⁴. Although laparotomy enabled direct visualization and aspiration of ovarian follicles, it was highly invasive, required extended postoperative recovery, and restricted the frequency of donor use. Consequently, this approach proved unsuitable for repeated collections or large-scale embryo production programs.

Subsequent advances led to the development of ultrasound-guided ovum pick-up (OPU) techniques, enabling minimally invasive oocyte collection and reducing surgical intervention^5,6,7,8. While this approach improved animal welfare and procedural efficiency, the relatively small body size and anatomical constraints of alpacas may limit probe positioning and follicle accessibility, potentially affecting consistency and recovery efficiency¹. Moreover, successful implementation requires specialized personnel training to ensure appropriate equipment handling, accurate follicular visualization, and reproducible outcomes⁷˒⁸.

Laparoscopic ovum pick-up (LOPU) has emerged as a significant advancement in assisted reproduction. Initially established in small ruminants^9.10.11, it has since been adapted across a broad range of species, including prepubertal cattle^12–14, deer¹⁵˒¹⁶, pigs¹⁷˒¹⁸, felids (e.g., pumas, jaguars, and tigers)¹⁹˒²⁰, and non-human primates²¹˒²², highlighting its versatility across domestic and wildlife species. The technique provides direct transabdominal access to the ovaries through minimally invasive entry, enabling precise follicular aspiration under real-time visual guidance⁹˒²³. Although LOPU has been extensively refined in prepubertal cattle^12–14, sheep¹⁰˒¹¹˒²³, and goats¹¹˒^24–27, its application in alpacas (Vicugna pacos) remains comparatively limited²⁸. Nevertheless, available evidence suggests that LOPU can achieve outcomes comparable to those reported in small ruminants while avoiding the need for rectal manipulation, supporting its potential as a technically robust approach for oocyte collection in this species²⁹˒³⁰.

Despite these advances, standardized and reproducible LOPU protocols for alpacas remain scarce. Most camelid ARTs studies have focused on in vitro maturation, fertilization, and embryo culture using oocytes derived from slaughterhouse ovaries or ultrasound-guided aspiration⁸˒³¹˒³². Although these approaches have provided valuable insights into oocyte competence and embryo development, they do not ensure a consistent supply of high-quality, in vivo–derived oocytes from genetically valuable donors. Furthermore, recent studies have highlighted the influence of the collection method on oocyte quality, underscoring the need for optimized in vivo retrieval techniques⁷.

The present study provides a detailed, step-by-step description of a laparoscopic ovum pick-up procedure specifically adapted for alpacas. This work is intended as a procedural and technical report rather than a comprehensive biological validation study. The protocol integrates current knowledge of camelid reproductive physiology, established principles from LOPU in small ruminants, and practical considerations related to anesthesia, ovarian superstimulation, laparoscopic access, and follicular aspiration. By presenting a reproducible and minimally invasive approach, this study aims to support the broader adoption of LOPU in alpaca reproductive programs and to facilitate future applications in in vitro embryo production, embryo transfer, and genetic engineering.

Protocol

Ethical Approval: All animal procedures were performed in accordance with institutional guidelines and approved by the Institutional Animal Care and Use Committee (IACUC) of Utah State University under protocol number 12895. All efforts were made to minimize animal suffering and to reduce the number of animals used.

The study was conducted at the Utah State University research facilities located in Logan, Utah, USA (41°44′ N, 111°50′ W; 1,381 m above sea level). The female alpacas were housed together in a single corral and provided ad libitum access to forage, a balanced commercial pelleted concentrate, mineral supplementation, and fresh water throughout the study period. To prevent uncontrolled mating and to eliminate potential male-induced behavioral and physiological effects on the female subjects, all intact males were housed in a separate barn located at a sufficient distance from the female corral. All animals were allowed an adequate acclimatization period prior to the initiation of experimental procedures. The research tools for this protocol are listed in the Table of Materials.

Number of animals used for LOPU:

Eight Huacaya alpacas (3–8 years of age) were selected as oocyte donors based on the following inclusion criteria: absence of reproductive tract abnormalities, adequate body condition score, and confirmed ovarian follicular response to superstimulatory treatment as assessed by transrectal ultrasonography prior to each procedure. Each animal underwent 10 repeated LOPU sessions at monthly intervals throughout the study period. All animals were maintained under uniform husbandry conditions for the duration of the study to minimize management-related sources of variation

A schematic overview of the laparoscopic oocyte collection procedure performed in live alpacas is provided in Figure 1.

Figure 1: Step-by-step representation of the LOPU protocol applied in female alpacas (Vicugna pacos) for oocyte recovery. The schematic depicts the four principal stages of the procedure: (1) cycle synchronization with GnRH administration, (2) follicular superstimulation with exogenous FSH, (3) laparoscopic aspiration of ovarian follicles, and (4) morphological evaluation and classification of recovered oocytes. This figure provides a visual summary of the methodological approach used in the present study. Please click here to view a larger version of this figure.

1. Animal restraint

1. To initiate restraint, firmly secure the head and neck of the alpaca against the chest of the handler while placing the opposite hand over the withers to maintain stability.
2. With assistance from a second technician, apply the Chukka technique³³ by placing a soft rope loop around the body immediately anterior to the pelvis, leaving approximately 15 cm of slack.
3. Gently elevate the hind limbs and position the hooves within the loop beneath the abdomen to provide stable support.
4. If the animal is in sternal recumbency (cushion position), one technician should gently elevate the posterior region of the body and slide the rope beneath the alpaca before positioning the hooves within the loop.
5. Once both hooves are properly positioned, tighten the loop securely and form a knot on the dorsal aspect of the animal to immobilize the hind limbs.
6. With the hind limbs immobilized and the alpaca maintained in the cushion position, transfer the animal to the diagnostic table with assistance from two technicians.
7. For additional stabilization, cross one forelimb over the head. For example, if the left forelimb is selected, position it on the right side of the head.
8. If the animal exhibits signs of nervousness, gently cover the eyes with a small towel to minimize visual stimuli and reduce stress.
9. Using hemostatic forceps, secure the tail by clipping it to the rump hair to prevent movement during the procedure.

2. Reproductive diagnostic and oocyte donor selection

1. Select donor animals based on the absence of reproductive abnormalities, appropriate nutritional management, good health status, normal growth records, and satisfactory reproductive performance.
2. Using a lubricated glove, gently evacuate the rectal ampulla, removing fecal material to minimize ultrasonographic artifacts.
3. Apply approximately 15–25 mL of sterile ultrasound gel to thoroughly lubricate the rectal ampulla, facilitating smooth probe insertion and reducing friction.
4. Gently introduce a 7.5 MHz transrectal ultrasound probe into the rectum.
5. Slowly and carefully manipulate the probe until the ovaries and associated reproductive structures are clearly visualized, avoiding excessive pressure or abrupt movements.
6. Record the number and diameter of follicles on each ovary. Particular attention should be given to follicles measuring ≥ 6.5 mm in diameter, which are classified as presumed dominant follicles (pDF).
   1. Animals presenting a pDF should be re-evaluated on the following day or within 48 h. If the follicle reaches a minimum diameter of 7.0 mm, it is classified as a true dominant follicle (DF).
   2. Animals exhibiting a confirmed DF are considered suitable candidates for inclusion as donors in superstimulation and oocyte collection protocols.
7. Acquire and store representative ultrasonographic images and measurements for documentation and subsequent analysis.
8. Upon completion of the examination, carefully withdraw the probe and allow the animal to stand and recover before returning it to the holding area.

3. Synchronization and superstimulation

1. Animals presenting a confirmed DF receive a single intramuscular (IM) injection of 50 µg gonadotropin-releasing hormone (GnRH).
   1. The day of GnRH injection is designated as Day zero (D0), marking the disruption of follicular dominance and the onset of follicular wave synchronization.
      CAUTION: GnRH is a synthetic peptide hormone; avoid accidental self-injection or mucosal exposure, as unintended administration may cause undesirable hormonal effects. Handle with gloves and follow institutional biosafety and handling guidelines.
2. At 48 h after GnRH administration (D2), ovarian superstimulation is initiated using a decreasing-dose regimen of follicle-stimulating hormone (Folltropin; NIH-pFSH-P1; FSH), administered intramuscularly in a twice-daily (AM/PM) schedule, with injections alternated between both sides of the animal, for a total dose of 200 mg.
   1. On D2, emergence of a new follicular wave is expected to coincide with the initiation of FSH treatment.
      CAUTION: FSH is a biological hormone preparation; handle with care to avoid accidental self-injection or mucosal exposure. Store according to manufacturer specifications and follow institutional biosafety protocols.
3. FSH treatments are administered from D2 to D5, and oocyte collection is performed on D6, approximately 12 h after the final FSH injection.
   NOTE: The complete synchronization and ovarian superstimulation protocol are summarized in Figure 2.

Figure 2: Timeline of the hormonal synchronization and ovarian superstimulation protocol used prior to LOPU in alpacas. A single intramuscular injection of 50 µg GnRH was administered on Day 0 (D0) upon confirmation of a dominant follicle (≥7 mm), followed by a decreasing-dose regimen of 200 mg FSH administered twice daily from Day 2 to Day 5 (D2–D5). Laparoscopic oocyte collection was performed on Day 6 (D6), approximately 12 h after the final FSH injection. Please click here to view a larger version of this figure.

4. Laparoscopic ovum pick-up

1. General Consideration
   1. To ensure procedural success and maintain high standards of animal welfare, all surgical procedures must be performed by a coordinated team
   2. Personnel involved in oocyte collection should be assigned clearly defined roles and adhere to established aseptic technique and ethical guidelines
      1. The primary surgeon leads the procedure and is responsible for anesthesia induction and monitoring, aseptic preparation, trocar and valve placement, ovarian manipulation, follicular aspiration guidance, wound closure, and postoperative care
      2. The assistant surgeon handles and positions the aspiration needle, assisting with follicular puncture and oocyte retrieval
      3. The camera operator manages endoscopic visualization, including camera positioning, focus adjustment, and lens cleaning to ensure optimal image quality throughout the procedure.
      4. Surgical support personnel are responsible for continuous monitoring of vital signs, CO₂ insufflation regulation, anesthetic administration, and documentation of physiological parameters during the procedure
2. Anesthesia
   1. Preoperative Fasting and Sedation
      1. Withhold hay, grain, and water for at least 36 h, 24 h, and 12 h, respectively, prior to anesthesia
      2. Sedate alpacas using a standard combination of ketamine, xylazine, and butorphanol (BKX cocktail) or equivalent anesthetic agents
         ​CAUTION: Ketamine, xylazine, and butorphanol are controlled substances and must be handled, stored, and administered strictly in accordance with institutional Drug Enforcement Administration (DEA) regulations and applicable federal guidelines
      3. Prepare the BKX cocktail by combining 10 mL ketamine (1,000 mg), 1 mL xylazine (100 mg/mL), and 1 mL butorphanol (10 mg/mL) to obtain a total volume of 12 mL.
      4. Inject at a dosing rate of 1 mL/18 kg IM, this corresponds to approximately 4.6 mg/kg ketamine, 0.46 mg/kg xylazine, and 0.046 mg/kg butorphanol.
   2. Anesthetic Induction and Intubation
      1. Approximately 10–15 min after BKX administration, induce anesthesia using isoflurane delivered via face mask.
      2. Secure the mask over the muzzle to minimize gas leakage and set the vaporizer to 5% isoflurane with an oxygen flow rate of 1–2 L/min.
      3. Once adequate anesthetic depth is achieved (e.g., absence of palpebral reflex), perform endotracheal intubation.
      4. Use a 43 French gauge (10 mm) internal diameter French endotracheal tube or smaller, as appropriate for individual animals.
      5. Insert a mouth gag (oral speculum) to maintain oral opening and visualize the larynx using a laryngoscope.
      6. Gently advance the endotracheal tube into the trachea using controlled rotational movements.
      7. Confirm correct placement by observing intermittent condensation at the tube opening during respiration.
      8. Inflate the cuff with approximately 20 mL of air to establish an adequate seal.
   3. Maintenance of Anesthesia
      1. Carefully remove the mouth gag and connect the endotracheal tube to the anesthesia circuit.
      2. Adjust the vaporizer to 1%–3% isoflurane with an oxygen flow rate of 1–2 L/min to initiate anesthetic maintenance.
      3. Continuously monitor respiratory function by observing the reservoir bag and inspiratory and expiratory valves of the anesthesia circuit.
3. Pre-Oocyte Collection
   1. Shave the abdominal region and remove any previously placed staples, if present. Thoroughly cleanse the area with surgical soap, disinfect with ethanol, and apply an iodine-based antiseptic solution.
   2. Transfer and secure the animal within the designated surgical area, ensuring proper alignment and maintenance of aseptic conditions.
4. Oocyte Collection
   1. Cover the abdominal region with a sterile surgical drape and create a fenestration to expose only the operative field
   2. Using a scalpel, perform a midline skin incision approximately 10–15 cm cranial to the udder
   3. Introduce a blunt insufflation needle into the abdominal cavity to establish capnoperitoneum using carbon dioxide (CO₂). Continue insufflation until adequate abdominal distension is achieved
      CAUTION: CO₂ used for abdominal insufflation is stored under high pressure and must be handled with care. Exposure to high concentrations of CO₂ in enclosed spaces may cause respiratory distress, dizziness, or asphyxiation. Ensure adequate ventilation in the surgical area, verify regulator and tubing integrity before use, and monitor insufflation pressure continuously throughout the procedure to prevent abdominal overdistension or gas embolism in the animal. Handle CO₂ cylinders according to institutional safety guidelines and applicable compressed gas regulations.
   4. Insert a 5 mm valved trocar through the initial incision to access the abdominal cavity and introduce the laparoscopic video camera.
   5. Place two additional 5 mm valved trocars bilaterally, positioned 10 cm lateral to the midline and 5 cm cranial to the udder, on the left and right sides of the first port.
      1. One port is used to introduce atraumatic Babcock forceps for ovarian manipulation and stabilization on the mesovarium, avoiding tissue injury.
      2. The second port is used for insertion of the aspiration needle.
   6. Position the female alpaca in the Trendelenburg position.
   7. Record the number and diameter of all visible ovarian follicles. Introduce the aspiration needle into the abdominal cavity and position it adjacent to the ovary.
   8. Using atraumatic forceps, gently manipulate and mobilize the ovary to optimize follicle exposure and release it from the infundibulum.
   9. Puncture each follicle and gently rotate the needle to ensure complete aspiration. Adjust the vacuum pressure to 25–30 mmHg and the flow rate to 13–15 mL/min, based on preliminary optimization and within ranges reported for oocyte recovery in small ruminants to minimize mechanical disruption of cumulus–oocyte complexes.
   10. Use a single-lumen aspiration system consisting of a 20-G short-bevel needle connected to a 50 cm cannula, silicone stopper, and 50 mL collection tube prefilled with collection medium.
   11. Generate negative pressure using an aspiration pump connected via silicone tubing.
   12. Upon completion of aspiration, rinse the ovarian surface with warm sterile saline solution containing heparin (0.1 mg/mL; 10 IU) to remove residual blood.
       NOTE: Heparin was added to the saline rinse medium (10 IU/mL) to reduce the likelihood of post-operative adhesion formation on the ovarian surface. The total dose administered was minimal and not expected to produce systemic anticoagulant effects.
       1. Using a 10 mL syringe introduced through the cannula, gently maneuver the ovary toward the cannula while simultaneously moving the cannula over the ovarian surface.
       2. Then flush and lightly agitate the ovary to ensure thorough blood removal and minimize adhesion formation.
   13. Close trocar incisions, use sutures or surgical staples, and perform simple interrupted skin sutures.
   14. Clean the surgical wounds with povidone–iodine and apply a topical repellent/healing ointment.
   15. For subsequent procedures, perform incisions lateral to previous surgical sites to minimize tissue trauma and reduce the risk of adhesion formation.
   16. At each LOPU session, evaluate ovarian tissue integrity, assess for adhesions, and inspect prior skin wound healing before proceeding.
       NOTE: Animals presenting with adhesions, compromised ovarian tissue, or inadequate wound healing should be removed from the donor group and excluded from further procedures.
   17. Administer 2 mL of cloprostenol intramuscularly (500 mcg/alpaca) if a corpus luteum is identified during the LOPU procedure.
       CAUTION: Cloprostenol is a synthetic prostaglandin analog; avoid skin contact and direct exposure, as it can be absorbed through the skin and may cause bronchospasm or uterine contractions. Handle with gloves and follow institutional safety guidelines.
   18. Administer ampicillin intramuscularly at a dose of 1.25 mL per 100 lb of body weight, for a final concentration of 8.8 mg/kg.
       ​CAUTION: Ampicillin is a broad-spectrum antibiotic; individuals with known penicillin hypersensitivity should avoid direct handling. Follow institutional guidelines for proper storage, administration, and disposal.
5. Post-Oocyte Collection
   1. At the conclusion of the procedure, reduce the vaporizer setting to 0% and maintain oxygen flow to facilitate anesthetic washout until the animal exhibits signs of recovery.
   2. Continuously monitor for pain and distress by assessing pulse oximetry values and body temperature using a temperature probe.
   3. Deflate the endotracheal tube cuff using a 20 mL syringe and carefully remove the endotracheal tube once protective reflexes return.
   4. Discontinue the oxygen supply following completion of anesthetic recovery.
   5. Allow animals to recover under continuous supervision until they regain sternal recumbency and are able to stand unassisted.
   6. Administer oral meloxicam at a dose of 15 mg per 100 lb of body weight for postoperative analgesia.
      CAUTION: Meloxicam is a non-steroidal anti-inflammatory drug (NSAID); prolonged or repeated skin contact may cause sensitization or irritation. Avoid ingestion, eye contact, and direct skin exposure. Handle with gloves and follow institutional safety and disposal guidelines.
   7. Once fully recovered and stable, return the animals to their respective stalls. The surgical procedure typically requires approximately 25–35 min to complete.
   8. Perform postoperative physical examinations, including assessment of body temperature, respiratory rate, and heart rate, twice daily for a minimum of 3 days.
   9. Additional monitoring includes evaluation of the surgical incision site, feed intake, urination, and defecation to ensure normal recovery and early detection of potential complications.

5. Oocyte recovery and morphological classification

1. Transfer the contents of each 50 mL conical collection tube (containing follicular fluid and collection medium) obtained during LOPU into a sterile Petri dish.
2. Examine each Petri dish under a stereomicroscope to identify cumulus–oocyte complexes (COCs).
3. Using a sterile pipette, recover oocytes from the medium and transfer them into maturation medium.
4. Classify oocytes according to morphological characteristics.
   1. Grade I oocytes are defined as those surrounded by at least two complete layers of compact cumulus cells and presenting a homogeneous, evenly granulated cytoplasm.
   2. Grade II oocytes are characterized by the presence of at least one intact layer of cumulus cells and a mostly homogeneous cytoplasm with minor irregularities.
   3. Grade III oocytes exhibit partial or incomplete layers of cumulus cells and may present cytoplasmic heterogeneity or irregular granulation.
   4. Grade IV oocytes lack cumulus cell investment and typically display a fragmented, degenerated, or clearly abnormal cytoplasm.

6. Descriptive statistics

1. Perform all descriptive statistical analyses using SAS software. Present data as mean ± standard deviation (SD).
2. Summarize follicular response, aspiration efficiency, and oocyte recovery parameters using standard descriptive statistics to evaluate the consistency and reproducibility of the protocol.

Results

Ovarian response and oocyte recovery following FSH superstimulation in alpacas are summarized in Table 1. Animals synchronized and superstimulated with 200 mg FSH developed an average of 14.88 ± 3.85 ovarian follicles per session, of which 10.94 ± 3.62 were successfully aspirated, resulting in a follicle aspiration rate of 74.18 ± 8.91%. A mean of 6.68 ± 2.21 oocytes were recovered per procedure, corresponding to an oocyte recovery rate of 61.30 ± 8.21%. Of the total oocytes collected, 59.05% were classified as Grade I–II, representing a substantial proportion of oocytes meeting morphological criteria for further use in in vitro maturation (IVM), in vitro fertilization (IVF), and in vitro culture (IVC) protocols.

Total Ovarian Follicles (n)	Ovarian Follicles Aspirated (n)	Follicle Aspiration Rate (%)	Oocytes Collected (n)	Oocyte Recovery Rate (%)	Oocyte Morphological Classification (%)
					G1	G2	G3	G4
14.88 ± 3.85	10.94 ± 3.62	74.18 ± 8.91	6.68 ± 2.21	61.30 ± 8.21	48.57	10.48	26.67	14.29
Values are presented as mean ± standard deviation SD.

Table 1: Ovarian response and oocyte recovery following FSH superstimulation and laparoscopic ovum pick-up (LOPU) in alpacas. Total ovarian follicles, ovarian follicles aspirated, and oocytes collected are expressed as averages across all collection sessions. Follicle aspiration rate, oocyte recovery rate, and oocyte morphological classification are expressed as percentages. Oocytes were morphologically classified as follows: Grade I (G1) — at least two complete compact cumulus cell layers and homogeneous cytoplasm; Grade II (G2) — at least one intact cumulus layer with mostly homogeneous cytoplasm; Grade III (G3) — partial or incomplete cumulus layers with cytoplasmic heterogeneity; and Grade IV (G4) — absent cumulus investment with fragmented or degenerated cytoplasm. Values are presented as mean ± standard deviation (SD), where applicable.

Representative laparoscopic images illustrating ovarian morphology and structures observed during LOPU are shown in Figure 3. Ovaries exhibiting a strong follicular response contained multiple well-developed follicles larger than 5–6 mm in diameter (Figure 3A–D). Following follicular aspiration, ovaries displayed reduced follicular turgidity and visible puncture sites corresponding to aspirated follicles (Figure 3E–H). In contrast, some animals exhibited limited follicular development, characterized by a small number of follicles and reduced ovarian activity (Figure 3I–L). Additional ovarian structures were occasionally observed during the procedure, including follicular cysts (FC) (Figure 3M, N). Finally, corpora lutea (CL) were observed as an indication of recent ovulation (Figure 3O, P).

Figure 3: Representative laparoscopic images of ovarian morphology and associated structures observed during LOPU in alpacas. Panels A–D illustrate a strong follicular response with multiple well-developed follicles (5–6 mm in diameter). Panels E–H show ovaries following follicular aspiration, with visible puncture sites and reduced follicular turgidity. Panels I–L depict limited follicular development representative of a poor ovarian response. Panels M–N show a follicular cyst (FC). Panel O shows a corpus luteum (CL). Panel P shows the right uterine horn and associated ovary. Scale bars = 5 mm. Please click here to view a larger version of this figure.

Descriptive analysis of follicular size distribution revealed a predominance of medium- to large-sized follicles at the time of collection (Figure 4). Follicles measuring 5–6 mm in diameter constituted a greater proportion of the ovarian follicular population compared with follicles measuring 3–4 mm. In contrast, the smaller follicle category (3–4 mm) exhibited greater variability and represented a lower percentage of the total follicular pool. These observations indicate that the superstimulation protocol promoted follicular development within the size range considered suitable for oocyte aspiration.

Figure 4: Ovarian follicle size distribution at Laparoscopic Ovum Pick-up (LOPU) in FSH-superstimulated alpacas. Box plots illustrate the percentage distribution of follicles within two diameter classes (3–4 mm and 5–6 mm) recovered during laparoscopic oocyte pick-up (LOPU) following GnRH synchronization and FSH superstimulation. The box represents the interquartile range (IQR; 25th–75th percentiles), the horizontal line within the box indicates the median, whiskers extend to minimum and maximum values, and the circle (○) denotes the group mean. Please click here to view a larger version of this figure.

Representative images of cumulus–oocyte complexes (COCs) and oocytes collected by laparoscopic ovum pick-up (LOPU) are shown in Figure 5. The micrographs illustrate variations in morphological quality, including well-preserved cumulus investment and homogeneous cytoplasm in high-grade oocytes, and reduced cumulus coverage and cytoplasmic heterogeneity in lower-grade oocytes.

Figure 5: Representative micrographs of immature cumulus–oocyte complexes (COCs) recovered from alpacas (Vicugna pacos) by laparoscopic ovum pick-up (LOPU) and classified according to morphological quality. Upper panels illustrate Grade I and II oocytes, characterized by compact cumulus cell investment and homogeneous cytoplasm. Lower panels show Grade III and IV oocytes, exhibiting partial or complete absence of cumulus cells and cytoplasmic irregularities. Scale bars = 100 µm. Please click here to view a larger version of this figure.

Collectively, the observed follicular response, aspiration efficiency, and oocyte recovery rates observed in the present study suggest that the hormonal synchronization and superstimulation regimen evaluated may support in vivo oocyte collection in alpacas. The proportion of aspiratable follicles and the recovery of morphologically suitable oocytes indicate that this protocol may be applicable to subsequent in vitro embryo production, embryo transfer, and related reproductive biotechnology procedures in this species, although studies with a larger number of animals are needed to strengthen and validate these findings.

Discussion

The present study indicates that the described synchronization and superstimulation protocol, combined with LOPU, supports ovarian response and efficient in vivo oocyte recovery in alpacas. Animals treated with 200 mg FSH developed a substantial follicular population, with approximately three-quarters of follicles successfully aspirated and a recovery rate exceeding 60%. All visible follicles greater than 2 mm in diameter that could be adequately stabilized were aspirated; however, a proportion of follicles remained inaccessible, likely due to insufficient firmness of ovarian fixation during the procedure, which may have limited complete needle access to all visible follicles. These outcomes nonetheless indicate that the hormonal regimen effectively induces follicular development while maintaining procedural efficiency during follicular aspiration.

GnRH administration at the time of dominant follicle detection has become the preferred synchronization approach in non-mated alpacas, given that during their prolonged follicular phase, dominant follicles repeatedly reach diameters sufficient to respond to exogenous GnRH, enabling reliable follicular wave turnover^34,35,36,37. While progesterone-based devices and prostaglandin protocols have been explored in Alpacas, these approaches have yielded inconsistent results due to device expulsion, vaginal complications, and unreliable follicular suppression³⁵, supporting the selection of the GnRH-based protocol used in the present study for its practicality and compatibility with LOPU for consistent oocyte recovery in alpacas.

The predominance of medium- to large-sized follicles (5–6 mm) at the time of collection is consistent with an active follicular response to the superstimulation protocol. In alpacas, follicle size has been associated with in vitro cytoplasmic maturation and developmental potential³⁸, suggesting that the follicular profile observed may favor the recovery of morphologically suitable cumulus–oocyte complexes (COCs), as illustrated in Figure 5. However, as the present study was not designed to directly compare FSH dosing regimens and the sample size was limited, future studies with larger animal numbers and comparative treatment designs would be needed to confirm whether the decreasing-dose FSH protocol specifically promotes superior follicular synchrony or oocyte quality in alpacas.

The morphological diversity observed among recovered oocytes reflects the inherent biological variability of superstimulation responses. High-grade oocytes exhibited compact, multilayered cumulus investment and homogeneous cytoplasm, features commonly associated with improved in vitro maturation and embryo development potential. In contrast, lower-grade oocytes displayed partial cumulus loss or cytoplasmic irregularities, which may compromise further developmental competence³⁹. Nonetheless, the overall proportion of recovered oocytes and the consistency of follicular response indicate that the protocol provides a reliable source of in vivo–derived COCs suitable for downstream applications.

Almost 60% of the recovered oocytes were classified as Grade I–II, representing a substantial proportion of morphologically suitable oocytes for in vitro procedures, particularly in settings where slaughterhouse-derived ovaries are unavailable or limited. In contrast, an important proportion of the recovered oocytes were classified as Grade III–IV, characterized by partial or complete absence of cumulus cell investment. This may be partially attributed to the aspiration conditions of the vacuum pump, including pressure settings, needle diameter, tubing length, and turbulence generated during follicular aspiration, as previously reported for oocyte collection procedures in other species²⁹. Optimization of these aspiration parameters may therefore represent an important consideration for improving oocyte morphological quality in future LOPU procedures in alpacas.

The aspiration flow rate represents a critical technical parameter during follicular oocyte recovery, as excessive pump speeds have been associated with mechanical disruption of the cumulus–oocyte complex (COCs) and a consequent reduction in oocyte morphological quality. With a constant-flow pump system to retrieve oocytes from ovine ovaries obtained from slaughterhouse, flow rates exceeding 20 mL/min were reported to be detrimental to oocyte morphological integrity, with an optimal range of 10–20 mL/min recommended to minimize aspiration-induced damage⁴¹ . In contrast, earlier LOPU studies conducted across various species have described aspiration rates of 50–70 drops per minute as operationally effective^{10,12,13,14,15,16} . More recently, an aspiration flow rate of 22 mL/min during OPU with transvaginal ultrasound-guided follicle aspiration was reported to result in marked morphological compromise of oocytes recovered in alpacas, further supporting the use of lower flow rates to preserve COCs integrity⁷. There is limited peer-reviewed literature on specific aspiration flow rate parameters for LOPU in alpacas.

In the present study, although a systematic comparison of aspiration flow rates was not conducted, a flow rate of 13–15 mL/min at a negative pressure of 30 mmHg was selected as the operative setting based on preliminary testing of multiple flow rate conditions. This range falls within the optimal interval described for sheep⁴¹ and is below the threshold associated with oocyte morphological deterioration in alpacas⁷. No apparent adverse effects on COCs morphology were detected under these conditions, suggesting that the operative parameters employed were appropriate. Future studies by systematically evaluating a broader range of aspiration flow rates under LOPU conditions in this species are warranted to establish evidence-based standardized protocols. LOPU has been demonstrated in small ruminants, including goats and sheep, to be a safe and repeatable procedure that can be performed multiple times in the same animal without inducing significant pain, postoperative complications, or detrimental effects on reproductive performance^23,40. However, in alpacas, the impact of repeated oocyte collection on animal welfare and subsequent reproductive efficiency has not been systematically evaluated. Although the present study was not specifically designed to assess these effects, it is worth noting that LOPU was performed 10 times on each animal over a 10-month period, and no adhesions or morphological alterations of the reproductive organs or abdominal wall were observed throughout the study. These observations are descriptive and were not derived from a study specifically designed to evaluate the long-term effects of repeated LOPU on reproductive performance or animal welfare. Systematic studies are nonetheless necessary to formally evaluate these variables in alpacas and determine whether repeated LOPU adversely affects short- and long-term reproductive performance under the conditions described.

Compared to ultrasound-guided ovum pick-up approaches, LOPU offers direct ovarian visualization and may enhance follicular access in alpacas, whose anatomical characteristics can limit the effectiveness of transvaginal techniques⁹. OPU in alpacas has been reported to achieve collection rates of approximately 75%, recovering a mean of 3.3 oocytes per ovary, with oocytes classified as Grade I and II representing only 30% of the total recovered⁷. In contrast, the LOPU protocol described in the present study yielded a collection rate of 61%, while the proportion of morphologically suitable oocytes classified as Grade I and II reached approximately 60% of total recovered oocytes, a substantially higher proportion than that reported with OPU.

Although the collection rate observed with LOPU was numerically lower than that reported for transvaginal OPU, the marked improvement in oocyte morphological quality suggests that LOPU may represent a more advantageous approach for downstream in vitro embryo production applications in alpacas. Both techniques share susceptibility to aspiration-related morphological compromise, likely attributable to vacuum pressure and flow rate conditions during follicular aspiration^7,8. Notably, oocyte recovery rates obtained with LOPU in alpacas are comparable to those reported in small ruminants^9,10,27, collectively demonstrating that laparoscopic retrieval is both feasible and reproducible in this species when appropriate hormonal preparation and technical expertise are applied. Several technical factors may influence oocyte harvest and quality outcomes, including follicular wave synchronization accuracy, timing of aspiration relative to the final FSH dose, surgeon experience, and aspiration system parameters such as vacuum pressure and flow rate, length of the aspiration line, tubing diameter, needle diameter, and the turbulence generated during the alternating suction and non-suction cycles of the pump⁴¹. Standardization of these parameters is essential to optimize oocyte harvest and morphological quality. Additionally, while morphological assessment provides a practical measure of oocyte evaluation, further studies incorporating in vitro maturation, fertilization, and embryo development endpoints would strengthen the functional validation of the protocol and confirm the developmental potential of oocytes recovered through LOPU in alpacas.

In conclusion, the combined application of GnRH-induced synchronization, decreasing-dose FSH superstimulation, and LOPU appeared to represent a feasible strategy for in vivo oocyte recovery in alpacas under the conditions described. The follicular response, aspiration efficiency, and oocyte recovery rates observed suggest that this protocol may be suitable for application in reproductive biotechnology programs in this species. No apparent adverse effects on ovarian integrity or donor health were observed across repeated procedures; however, systematic evaluation of the long-term impact of repeated LOPU sessions on reproductive performance in alpacas is still needed. These findings may contribute to the growing body of evidence supporting the application of assisted reproductive technologies in South American camelids and could serve as a preliminary foundation for future investigations into in vitro embryo production, embryo transfer, and related germline-based procedures in alpacas, provided that studies incorporating larger animal numbers are conducted to generate more robust and generalizable data.

Limitations

The present study has limitations that should be acknowledged. First, the relatively small number of donor animals limits the statistical power of the findings and may not fully capture the biological variability inherent to the alpaca population. Second, oocyte quality was assessed solely by morphological classification, without functional validation of developmental competence via IVM, IVF, and IVC. Morphological grading, while practical and widely used, does not necessarily reflect the true developmental potential of recovered oocytes. Future studies incorporating these in vitro endpoints are therefore essential to further evaluate the protocol and to confirm the functional suitability of LOPU-derived oocytes for embryo production and advanced reproductive biotechnology applications in alpacas.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Ampicillin	Valleyvet	1787RX	Ampicillin Powder for Injectable Suspension 25 mg
Aspiration Needle	WTA Technologies	26041	20G
Aspiration Pump for bovine OPU	Minitube	23362/0001	Warming block, foot pedal, 115 V
Babcock Forceps	Karl Storz	26166	5 mm, 30 cm
Blunt Cannula	Medline	MDS0775641	23 G
Camerea Head	ESC Medicams	ESC2000212	HD, portable endoscopy camera
Carbon Dioxie	Airgas	N/A	20-pound cylinder, USP Medical, CGA 320 washer
Chlorheadine 2% solution	Durvet	7-45801-10259-0	N/A
Cradle for laparoscope	Custom-built at Utah State University	N/A	Inclination range of 30–45°, with extended support attachment for the alpaca neck
Disposable Scalpel Blades	Medline	CISION22SS	Stainless-Steel Blade, No. 22. Sterile
Disposable Skin Stapler	Medline	QTX253001H	35 mm
Dulbecco's Phospate Buffered Saline	Gibco	14190144	No Calcium, No Magnesium
Endotracheal tube	Penn Veterinary Supply, Inc	JORJ615-H	43Fr 10.0 mm silicone cuffed
Estroplan	AmerisourceBergen	52077	Cloprostenol 250 mg/mL
Fertagyl	AmerisourceBergen	60654	Gonadorelin 43mg/mL
Fiber-Optic Light Cable	Olympus	WA03210A	5 mm
Folltropin Dual Pack	Professional Embryo Transfer Supply, Inc	04-020-129-3	Porcine Pituitary-Derived Follicle Stimulating Hormone 700 IU FSH
Halogen Ligh Source	Olympus	CLC-SC	Halogen Lamp, 120 V, 1.7A
Heparin Sodium Salt	Sigma-Aldrich	H4784	≥180 USP units/mg
Insuffation Tubing	Medline	DYNJ05933	Luer, Sterile
Isoflurane vaporaizer	VetEquip	911103	Equipped with scavenging collection bag
Ketamine	Dechra	1000001250	Ketamine Hydrochloride 100 mg/mL
Lubricant	Durvet	7-45801-11406-7	All purpose
Medium 199	Gibco	11150059	Earle's Salt, No suplemented
Meloxicam	Valleyvet	1545RX	Meloxicam tablets,  USP 15 mg
Monitor series S32GF	Samsung	‎LS24F320GANXZA	FHD, 24 inches
Rigid Laparoscope	Olympus	A50372A	5 mm, 0°, 30 cm
Rompun	Dechra	1000001151	Xylazine 100 mg/mL
SAS software	SAS Institute Inc.	N/A	Used for descriptive statistical analysis
Scalpel Handle	Medline	MDS10801	#3
Stereo Microscope	Olympus	SZ61	Transmitted light mirrored base
Sterile Disposable Drapes	Medline	LPC980927	N/A
Torphadine	Dechra	1000001776	Butorphanol Tartrate 10 mg/mL
Trocar-Cannula	Endoscopy Superstore	101021-A	5 mm, Spiral, with automatic valve
Ultrasound Diagnostic System	Chison	Eco1	Lineal Probe 7.5 MHz