Embryo Transfer Surgery via Laparotomy in Gilts

Summary

This protocol describes the application of the surgical technique used to transfer cloned pig embryos via laparotomy in gilts.

Abstract

This protocol aims to demonstrate the surgical technique for transferring cloned pig embryos to the oviduct, a method widely used in the production of genetically modified pigs for biomedical research. Nine gilts underwent hormonal synchronization and laparotomy for the transfer of cloned embryos produced by somatic cell nuclear transfer (SCNT) at stages of up to 4 cells on day 2 to the oviduct. Gestational diagnosis was conducted via ultrasound examination 30 days post-transfer surgery. Six out of the nine operated gilts exhibited signs of pregnancy on ultrasound examination. However, as there was no progression in fetal development as assessed by echography, the gilts underwent necropsy at 60 days for the collection of biological material and assessment of the reproductive system. Adhesions were observed in the uterine horns, ovaries, and oviducts. From the uterine lumen of two of the euthanized gilts, one and four embryonic structures with gestational ages ranging between 12 to 20 days were obtained. Despite the absence of live piglets, likely attributed to the low-efficiency rate of transferring cloned pig embryos, which is influenced by various factors, including the number and quality of transferred embryos, the presented surgical technique proved to be rapid and safe.

Introduction

Pigs are an excellent experimental model for biomedical research due to their anatomical, physiological, and genetic similarities to humans¹. These animals have often been used in research related to xenotransplantation, with the intention of producing organs, cells, or tissues that promote a low risk of rejection when transplanted into humans. Xenotransplantation research aims to increase the organ supply for human transplantation, thus reducing the waiting list of patients².

The production of pigs for xenotransplantation involves several steps, including the production of clones from genetically edited swine cells. After the in vitro production of genetically modified cloned embryos, the embryos are transferred to the reproductive system of a sow with a synchronized estrous cycle to prepare the uterine physiology for the reception and gestation of the new conceptus³.

Embryo transfer in pigs can be performed by non-invasive or invasive methods⁴. Among the non-invasive methods is transcervical transfer, which does not require any surgical intervention. However, this method is restricted to transferring embryos at later development stages (i.e., morula or blastocyst stages) and does not allow for the precise determination of the catheter insertion site or embryo deposition⁵. Laparoscopy and laparotomy are considered invasive methods of embryo transfer. Laparoscopy is less invasive but requires specific and costly equipment, and its efficiency varies considerably (from less than 20% to over 80%) due to various factors such as difficulty in manipulating reproductive structures and the type of catheter used^4,6. Therefore, transfer via laparoscopy is still less efficient compared to surgical transfer methods via laparotomy⁴.

Embryo transfer surgery in sows via laparotomy is a relatively simple and quick procedure, typically taking about 30 min. However, it should be performed in a surgical center equipped with inhalation anesthesia apparatus and a specialized team. For commercial pig strains (such as Landrace, Large White, or their crossbreeds), special equipment like hoists for lifting gilts and a wide, sturdy surgical table are necessary due to the animals' considerable weight (around 130-150 kg).

For the surgery to be successful, the embryos must be evaluated for their stage of development beforehand. Embryos of up to 4 cells are recommended to be transferred into the uterine tube. Embryos at stages beyond 4 cells, such as morulas and blastocysts, should be transferred to the uterine horn^7,8.

Although research groups worldwide are performing the production and surgical transfer of genetically modified cloned pig embryos, there are still no well-defined protocols demonstrating this procedure through videos. This approach is crucial for the success of gestation, as the technique requires precise deposition of embryos in the exact location of the oviduct, involving the localization of the tubal ostium and the introduction of the pipette containing the embryos. This technique can be better understood through explanatory videos of the entire procedure. Therefore, this article aims to demonstrate the laparotomy surgery for transferring cloned embryos to the oviduct of gilts, an essential prerequisite for the future production of genetically modified pigs to be used for xenotransplantation or other related purposes.

Protocol

This study was approved by the Ethics Committee on Animal Use in Research of the College of Veterinary Medicine and Animal Science of the University of São Paulo, protocol number 6088030523. Nine seven-month-old gilts from the Água Branca nucleus pig farming, located in Itu city, São Paulo state, Brazil, were used right after the second estrus detection⁹. The details of the reagents and the equipment used in the study are listed in the Table of Materials.

1. Animal preparation

1. Fast the animals for 12 h and restrict water access for 4 h before the surgical procedure.
2. Using a 40 mm x 12 mm needle, draw up 10 mg/kg of ketamine and 0.2 mg/kg of midazolam in the same 50 mL syringe. Attach the syringe to an IV administration set to facilitate medication administration, maintaining a safe distance from the animal and allowing it to move freely.
3. Administer the pre-anesthetic medication via intramuscular injection into the trapezius muscle, caudal to the external ear canal.
4. Cannulate the auricular vein with a 20 G catheter and connect it to a macrodrip administration set for intraoperative fluid therapy with 0.9% NaCl solution at a rate of 10 mL/kg/h.
5. Induce anesthesia with 4 mg/kg of propofol intravenously (following institutionally approved protocols).
6. Perform orotracheal intubation of the animal using a laryngoscope and a size 9 orotracheal tube with a cuff¹⁰.
7. Maintain anesthesia with 1.5% to 2% isoflurane inhalation. Additionally, monitor anesthesia by evaluating physiological parameters such as heart rate, which should be maintained between 100 and 120 beats per minute, oxygen saturation, which should be kept between 99% and 100%, muscle relaxation, absence of movement, and reflexes evaluation.
   NOTE: Body temperature should also be monitored and controlled, staying between 37.5-38.5 °C. When necessary, in addition to isoflurane, propofol can be used to maintain control of the parameters. Fentanyl hydrochloride should be administered as an analgesic at a dose of 0.02 mg/kg every 20 min. All parameters should be recorded every 5 min.
8. Administer perioperative analgesia with a single dose of 0.002 mg/kg of fentanyl intravenously and 4 mg/kg of tramadol hydrochloride intramuscularly into the gluteal or trapezius muscle.
9. Place cardiac electrodes on the thoracic region for intraoperative monitoring of heart rate. Record heart rate, rectal temperature, and oxygen saturation every 5 min.

2. Surgical procedure

1. Position the gilt on the surgical table in dorsal recumbency.
2. Cover the animal's legs with procedural gloves to minimize contamination of the surgical field.
3. Perform ventral caudal abdominal hair removal between the penultimate pair of mammary glands using a surgical clipper.
4. Perform preoperative skin antisepsis with a chlorhexidine-impregnated antiseptic brush, making circular movements from the umbilical scar region to the last pair of mammary glands. Remove chlorhexidine with a clean, moistened compress.
5. Ensure that the surgeon and the assistant perform surgical antisepsis and wear sterile surgical gowns and gloves for surgery.
   NOTE: The animal should be prepared for surgery in a preparation area separate from the surgical site.
6. Prepare the surgical table with the appropriate and previously sterilized instruments listed in the Table of Materials.
7. Perform final surgical field antisepsis using sterile gauze and chlorhexidine antiseptic, cleaning the region in a fishbone pattern, starting at the incision site and expanding laterally.
8. Repeat this procedure three times with chlorhexidine antiseptic, and then repeat the procedure with alcoholic chlorhexidine.
9. Completely cover the animal with a sterile surgical drape, and using scissors, create a rectangular opening in the surgical drape, approximately 10 x 20 cm², if the drape is not fenestrated.
10. Position the surgical drape over the penultimate pair of inguinal mammary glands. Place 4 Backhaus clamps to secure the surgical drape over the animal's skin.
11. Incise the skin with a scalpel, creating an incision approximately 10 cm in length.
12. Achieve hemostasis of subcutaneous vessels with hemostatic forceps and/or absorbable suture size 2.
13. Dissect the subcutaneous tissue with fingers or with the aid of fine-pointed scissors, deepening the incision to reach the linea alba in the abdominal musculature.
    NOTE: Palpate the midline of the abdomen to feel the linea alba as a more rigid and fibrous area; observe the separation of the rectus abdominis muscles and note the change in tissue consistency while dissecting to identify the linea alba accurately¹¹.
14. With Allis forceps, grasp the musculature and lift it to make a stab incision in the linea alba with a scalpel.
15. Insert the index finger into the incision to assess the presence of adhesions. Extend the incision over the linea alba caudally and cranially with blunt/blunt or fine/blunt scissors carefully to avoid injuring other organs and underlying structures.
16. Make an incision of sufficient length so that the surgeon's one hand can be inserted into the abdominal cavity to locate and exteriorize the uterine horns and/or ovaries. A Gosset retractor may be used to facilitate entry into the abdominal cavity and suturing of the musculature in a single layer at the end of the procedure.
17. If one of the uterine horns is located before the ovaries, gently follow it until reaching the tip of the uterine horn and the ipsilateral ovary. During this procedure, assess the presence of fluid inside the uterus, which may compromise gestation.
18. After locating the ovary, gently pull it and carefully expose it. Assess the presence of pre-ovulatory follicles, hemorrhagic corpora lutea, cyclic corpora lutea, and corpus albicans to evaluate if the gilt's synchronization is as expected.
19. Gently remove the fimbriae of the uterine tube covering the ovary, evert its mucosa, and locate the ostium of the uterine tube. 
    NOTE: If the surgeon finds it difficult to identify the ostium of the fimbriae, a pocket on the ampulla of the oviduct can be done with a round-bodied suture needle, avoiding puncturing blood vessels, to insert the Tomcat catheter containing the embryos.
20. Carefully insert a Tomcat catheter containing the embryos into the ostium of the fimbriae until reaching the ampulla region of the uterine tube.
21. Attach a 1 mL syringe to the Tomcat catheter to push the embryo-containing fluid into the uterine tube. Cover the ovary with the fimbriae and return it to the abdominal cavity.
22. Locate the contralateral uterine horn and ovary and repeat steps 2.17-2.21.
23. Reposition the structures inside the abdominal cavity and add 1 L of saline solution or Ringer's lactate solution warmed (39 °C) to prevent adhesions before initiating abdominorraphy.
24. Suture the abdominal musculature with absorbable suture size 2 in "X" stitches (Sultan) interrupted.
25. Approximate the subcutaneous tissue with a continuous mattress suture using absorbable suture size 2.
26. Close the skin with size 2 nylon suture in interrupted simple stitches, surgical staples, or ethyl-cyanoacrylate adhesive. In this procedure, we opted for the use of adhesive.
27. Clean the surgical wound with gauze soaked in hydrogen peroxide and dry with a compress to enhance the fixation of the surgical adhesive on the surgical wound. Apply rifamycin or Pearson's ointment, cover with gauze, and finally, apply surgical adhesive.
    NOTE: The wound should remain closed with a dressing for 5 days.

3. Postoperative care

1. Evaluate the animals for signs of characteristic clinical pain for at least the next 3 consecutive days, according to the scale proposed by Luna et al.¹².
   NOTE: This scale is based on different behaviors exhibited by pigs when experiencing different levels of pain, such as posture, interaction and interest in the environment, activity, appetite, and attention to the affected area. The score ranges from 0 to 3, depending on the type of behavior exhibited, for each parameter evaluated. At the end of the evaluation, the sum of all scores reflects the intensity of pain experienced by the animal, where 0 represents no pain, and 18 represents very intense pain¹².
2. Administer a single dose of 15 mg/kg of amoxicillin and 0.4 mg/kg of 2% meloxicam both intramuscularly (in the gluteal or cervical muscle) once a day for 3 days as postoperative care to prevent pain, inflammatory processes, and infection.
   NOTE: If the pain score is equal to or greater than 6 on the pain scale used (score from 0 to 18), administer 5,000 mg/animal of sodium dipyrone intramuscularly (in the gluteal or cervical muscle) once a day in addition to the anti-inflammatory and antibiotic. Alternatively, perform analgesia as outlined in the local IACUC guidelines.
3. On the 6th day after surgery, remove the surgical adhesive for wound cleaning with gauze soaked in 2% alcoholic chlorhexidine solution and apply topical Pearson's ointment until the 10th day, when suture removal or surgical staples removal, if used, is recommended.

4. Ultrasonographic gestational diagnosis of pregnancy

1. Perform transabdominal pregnancy ultrasonographic evaluation with an ultrasound device (frequency range 4.5 to 8.5 MHz) 30 days after embryo transfer surgery.
2. Apply ultrasound gel to the probe and position it near the inguinal abdominal region, facing medially.
   NOTE: Visualization of rounded anechoic embryonic vesicles, due to fluid in the uterine lumen, indicates gestation. Non-visualization of intrauterine fluid, or presence of intrauterine fluid without delineation of embryonic vesicles, suggests the absence of gestation.

5. Euthanasia of the animals

NOTE:  Euthanize the gilts that did not present fetuses of size and appearance corresponding to gestational age on the ultrasonographic examination.

1. Using a 40 mm x 12 mm needle, draw up 15 mg/kg of ketamine and 2 mg/kg of midazolam in the same 50 mL syringe attached to an intravenous administration set, and administer intramuscularly into the trapezius muscle in the cervical region caudal to the external ear canal, as preanesthetic medication.
2. Cannulate the auricular vein with a 20 G catheter and induce anesthesia with 4 mg/kg of propofol intravenously in a bolus until bradycardia occurs (following institutionally approved protocols).
3. Subsequently, administer 10 to 20 mL of 19.1% potassium chloride intravenously. Confirm cardiopulmonary arrest by observing the absence of respiratory movements, pale mucous membranes, loss of corneal reflex, and absence of heartbeats by placing a stethoscope over the cardiac region.
4. Collect any type of intrauterine content (embryonic tissue, liquid, or mucous secretions) from necropsied females¹³ and forward it for microbiological analysis to detect possible pathogens and also for genotypic characterization of the clones (if needed). Visually check the presence or absence of corpora lutea, ovarian cysts, abnormal secretions, and the occurrence of adhesions in the uterus, ovaries, and oviducts¹⁴.

Results

This article aims to demonstrate the laparotomy surgery for transferring cloned embryos to the oviduct of gilts. All animals remained in an adequate anesthetic plane, without any intraoperative incidents or complications during anesthesia recovery. The gilts took, on average, 2-3 h to stand up after the surgery ended.

All surgical procedures lasted, on average, 44 min. Nine gilts underwent surgery, with an average of 185 cloned embryos transferred per gilt, totaling 1,664 embryos transferred bilaterally (Table 1). The localization of the ovaries, their exteriorization, and the introduction of the Tomcat catheter containing the embryos into the ostium of the oviduct corresponded to the most delicate moments of the surgery. The ovaries of all gilts were inspected during surgery, and all presented recently ovulated hemorrhagic follicles, consistent with what was expected for the estrus synchronization protocol performed, which aimed to carry out embryo transfers shortly after ovulation occurrences. All procedures occurred without intraoperative incidents.

No animal showed signs of intense pain after surgery, with an average pain score of 0.33, 0.33, 3.67, 1.33, 0.33, 1.33, 4.67, 2.00, and 0.33 on the pain scale (ranging from 0 to 18) performed over the 3 postoperative days for each of the operated gilts (Table 2). Only 1 of the operated gilts presented subcutaneous fluid accumulation (seroma) on the 12^th day post-surgery, with signs of infection and inflammation localized near the surgical wound. After draining the fluid with a 40 mm x 1.6 mm needle guided by ultrasound, applying cold compresses to the inflamed area for 30 minutes daily for 7 days, administering ceftiofur (3 intramuscular injections, with a 72 h interval between injections) and 2% meloxicam (0.4 mg/kg SID, intramuscularly, for 3 days), the wound fistulated and drained the remaining mucopurulent content, resulting in complete improvement of the animal. The other operated gilts remained with the surgical dressing for 5 days without the need to change or remove it for wound cleaning, which was only performed from the 6^th to the 10^th day post-surgery.

In gilts 2, 8, and 9, no fluid accumulation and embryonic vesicle formation were observed in the uterine lumen (Figure 1A). Therefore, the gestational diagnosis of these animals was considered negative. The other gilts showed intrauterine fluid accumulation and the formation of embryonic vesicles visualized as well-defined anechoic rounded structures (Figure 1B-D). It was possible to identify, in two of the recipient gilts, a hyperechoic region relative to the embryonic structure in the early development phase inside the embryonic vesicles on the 32^nd day of gestation (Figure 1E,F).

After a negative gestational diagnosis or non-progression of gestation, the gilts were euthanized, and the reproductive system was analyzed. Adhesions in regions of ovaries, oviducts, and uterine horns to a lesser or greater degree were observed in gilts 1, 2, 3, and 4, and ovarian cysts were observed in gilt 8 (Figure 2). Four embryonic structures, with a gestational age compatible with approximately 12 to 20 days of gestation, were obtained from the uterus of gilt 3, and one structure was obtained from gilt 7 (Figure 3). The analysis of embryonic tissue fragments resulted in negative for the detection of Parvovirus, Leptospira sp., Erysipelothrix rhusiopathiae, and Porcine Circovirus types 2 and 3 (PCV2/PCV3), which can cause embryonic death or abortions.

Figure 1: Ultrasonographic images of the uterus of embryo recipient gilts 30 days after the surgical procedure. (A) The uterus of gilt 2 without the presence of fluid inside, characterizing a negative gestational diagnosis. The gravid uterus of gilt 1 (B), gilt 3 (C), and gilt 4 (D) containing embryonic vesicles with fluid inside (asterisks). The gravid uterus of gilt 3 (E) and gilt 7 (F) showing a hyperechoic structure (arrow) inside the embryonic vesicle, indicating fetal formation in the early phase at 32 days of gestation. Please click here to view a larger version of this figure.

Figure 2: Analysis of the reproductive system. Images of uteri, ovaries, and oviducts after the necropsy of gilt 1 (A), gilt 2 (B), gilt 3 (C), gilt 4 (D), gilt 5 (E), gilt 6 (F), gilt 7 (G), gilt 8 (H) and gilt 9 (I). Arrows highlight adhesions in uterine horns, ovaries, and oviducts, and arrowheads highlight ovarian cysts. Please click here to view a larger version of this figure.

Figure 3: Gestational diagnosis. Early-stage embryos removed from gilt 3 (A-D) and gilt 7 (E) present a positive gestational diagnosis and hyperechoic structures inside embryonic vesicles on ultrasonographic examination. Please click here to view a larger version of this figure.

Table 1: Number and type of embryos transferred per gilt, diagnosed pregnancies, and number of embryos obtained after necropsy of the operated gilts. Please click here to download this Table.

Table 2: Values assigned to each of the operated gilts regarding the presentation of pain signs. Please click here to download this Table.

Discussion

The surgical method described has been previously performed by other research groups working with the production of cloned pigs or genetically modified cloned pigs, with reports of births after the implementation of this technique^{15,16,17,18,19,20}. The pregnancy and birth rates of SCNT embryos produced from various cell lines range around 20%-30% (out of 264 transfers since 2017), with an average of 3.2 ± 0.4 piglets born per litter^3,21. Several factors are related to the low efficiency of this technique, including the synchronization protocol used for recipient sows, synchronization between the recipient sow and the stage of transferred embryos, the site of embryo deposition, the quality of oocytes used for SCNT and produced embryos, the quality and cellular stage of nucleus donor somatic cells, and the number of embryos transferred²¹.

According to the present study, the described surgery proved to be effective, fast, and safe for performing surgical transfer of cloned embryos (up to 4-cell stage) in gilts, resulting in positive ultrasonographic gestational diagnoses and visualization of embryonic structures inside the uterine lumen after necropsy, with gestational age approximately between 12-20 days of gestation²². Another less invasive method for transferring SCNT embryos to the oviduct of sows includes laparoscopy. However, the efficiency of this technique varies considerably from less than 20% to over 80% due to various factors such as the anatomy of the structures, the type of catheter used, and the site of embryo deposition^{4,6,23,24,25,26,27}. When transferring 2-4 blastomere pig embryos by laparoscopy via the infundibulum to 6 sows, Wieczorek et al.⁴ did not achieve positive pregnancies at 28-31 days after the procedure, and this result was related to difficulties in inserting the catheter into the oviduct, shallow catheter insertion into the oviduct, and occurrence of infundibulum perforation with the catheter. Therefore, the method of embryo transfer via laparotomy to the oviduct of sows still seems to be the most efficient.

The surgical risk associated with the laparotomy procedure for embryo transfer in sows is small; however, it is important to note the possibility of malignant hyperthermia during anesthesia, which is commonly seen in commercial pigs of the Landrace and Large White breeds²⁸. In this case, none of the operated gilts showed clinical signs of malignant hyperthermia during anesthesia. There is also a risk of minor local bleeding from larger vessels in the subcutaneous region, which can be easily stopped with hemostatic forceps or suturing with absorbable suture. The possibility of accidentally incising any organ of the abdominal cavity, such as the bladder or intestinal loops, during the incision with the scalpel in stab and extension incision in the linea alba with scissors may also occur. In this case, the surgeon must pay close attention during the incision of the linea alba and suturing of the abdominal muscles to avoid cutting or lacerating underlying structures.

The surgeon's manual dexterity is essential for proper manipulation of the ovaries and careful exteriorization, avoiding rupture of the vessels of the ovarian pedicle. When there is difficulty in locating the ovary(s), it is possible to start the procedure by locating one of the uterine horns and following its extension until reaching the tip of the uterine horn at the utero-tubal junction near the ovary. Care should be taken to avoid ovarian and uterine adhesions due to excessive manipulation of these organs. Hydrating these structures with lactated Ringer's solution or saline solution at 39 °C during surgery can help prevent adhesion formation²⁹. Manipulation of the uterine tube, especially in the region of the fimbriae, should be performed very delicately to avoid stretching and rupture. Although unilateral embryo transfer, only to one of the uterine tubes, can be performed to avoid excessive manipulation of the uterus and reduce surgical time, bilateral transfer may be safer and more advantageous, especially in cases where there is an accidental rupture of one of the uterine tubes or doubts about embryo deposition in the correct location. Furthermore, bilateral transfer is more recommended as it promotes higher pregnancy rates and number of piglets born when compared to unilateral transfer^16,30. The presence of cysts is associated with regular or irregular return to estrus, with up to 10% of females with ovarian cysts remaining in anestrus. It is related to various causes such as stress, use of hormonal protocols, or genetic inheritance^31,32. In our case, follicular cysts occurred despite following the dose and administration time recommended by the hormone manufacturer for heat synchronization.

Knowing how to locate the ostium of the uterine tube by the opening of the fimbriae is of fundamental importance for the correct deposition of embryos in the appropriate location. To avoid mistakes regarding the location of embryo deposition, it is recommended that prior training be performed on the passage of the Tomcat catheter filled with solution (e.g., water) through the ostium of the uterine tube in slaughterhouse specimens. When the pipette is introduced into the correct location, it is possible to observe an increase in the volume of the uterine tube upon injecting the solution. It is worth noting that the smaller the amount of fluid transferred along with the embryos, the better the chances of pregnancy³³. If the surgeon finds it very difficult to find the ostium of the uterine tube, it is possible to make a pocket on the oviduct and insert the catheter through it. Although this last method is faster and involves less manipulation of the oviduct, the surgeon needs to avoid blood vessels and be sure that the catheter has been inserted correctly inside the lumen of the oviduct.

Although the risk of postoperative infections from this surgical procedure is low, it can occur due to factors such as the cleanliness level of the area where the animals are housed, the duration of the surgical procedure, the maintenance of the dressing for an adequate time, and the correct prescription of medications, such as preventive antibiotic therapy and anti-inflammatories. The most plausible hypothesis for the occurrence of the infection observed in one of the operated gilts is related to the dressing falling off shortly after the animal gets up after anesthesia. Although daily cleaning of the surgical wound of this gilt was performed with gauze soaked in saline solution and topical application of Pearson's ointment, such a procedure was not enough to prevent ascending contamination, requiring cold water application at the site, drainage, systemic antibiotic therapy, and an additional 3 days of anti-inflammatory treatment. Even with the occurrence of subcutaneous infection, this gilt showed signs of pregnancy on ultrasonographic examination.

After using nylon threads and surgical staples to close the skin in previous surgeries, we chose to use ethyl-cyanoacrylate adhesive to reduce the stress when removing the stitches and prevent any remaining threads or staples. The presence of threads or staples promotes small local inflammation with pustule formation. The ethyl-cyanoacrylate adhesive has already been used safely in surgical procedures, proving to be effective, non-toxic, and bactericidal^34,35. All animals showed excellent wound healing, without complications, with the exception of the gilt, which had the dressing fall off shortly after the surgical procedure.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.25 mL straw	generic	-	Surgical material
1 mL syringe	Descarpack	341001	Surgical material
10 mL syringe	Descarpack	324601	Surgical material
20 mL syringe	Descarpack	324801	Surgical material
3 mL syringe	Descarpack	324201	Surgical material
5 mL syringe	Descarpack	324401	Surgical material
60 mL syringe	Descarpack	323201	Surgical material
9 mm endotracheal tube	Rusch	112482-000090	Surgical material
Allis forceps	generic	-	Surgical instrument
Amox LA	JA Saúde Animal	MAPA registration: 8.781/2004	Pharmaceutical drug
Bakhaus forceps	generic	-	Surgical instrument
Catheter 20G	Descarpack	362401	Catheter for intravenous access
Cetamin	Agener União	MAPA registration: SP-000292-5.000011	Anesthetic
Conductive clinical gel	RMC	ANVISA registration: 80122200013	Surgical material
Dipyrone D-500	Zoetis	MAPA registration: SP0000728-46	Pharmaceutical drug
Disposable scalpel n. 22	Wiltex	ANVISA registration: 10150470565	Surgical instrument
Disposable sterile sponge-brush	Rioquimica	7.89778E+12	Surgical asepsis
Easy-Scan:Go	IMV	ESCG01	Ultrasound
Endozime AW Plus	Ruhof	34514	Detergent for surgical instruments
Fentanil (Fentanest)	Cristália	ANVISA registration: 1029800810159	Anesthetic
Gosset retractor	generic	-	Surgical instrument
Halstead-mosquito hemostatic forceps	generic	-	Surgical instrument
Healing ointment - Unguento Pearson	Pearson SA	MAPA registration: SP0000094-16	Pharmaceutical drug
hydrogen peroxide solution	Rioquimica	ANVISA registration: 218690015	Surgical material
Isofluorane	Biochimico	ANVISA registration: 100630222	Anesthetic
IV Drip set extensor	generic	-	Fluid therapy
IV Macro drip set	Descarpack	410301	Fluid therapy
Lactofur (ceftiofur)	Ourofino SA	MAPA registration: SP0000051-50	Pharmaceutical drug
Laringoscope	-	-	Surgical instrument
Maxicam 2%	Ourofino SA	MAPA registration: SP0000051-69	Pharmaceutical drug
Micropore adhesive 5 cm x 10 cm	generic	1530	Surgical material
Midazolam	Hipolabor	ANVISA registration: 1134301430035	Anesthetic
Multi-Way IV Infusion Set	Descarpack	413201	Fluid therapy
Needle 40 mm x 1.2 mm	Descarpack	353601	Sterile needle for applying medicines and anesthetics.
Needle 40 mm x 1.6 mm	Wiltex	ANVISA registration: 10150470664	Sterile needle for applying medicines and anesthetics.
Needle holder	generic	-	Surgical instrument
Nylon 2 suture trhead	Shalon Medical	N502CTI40	Surgical material
Ordinary pen	generic	-	Regular pen for taking notes
Physiological solution 0.9% 500 mL bag	JP Farma	MS:1.0491.0070	Fluid therapy
Polyglycolic acid 2 suture thread	Atramat	G4099-75H	Surgical material
Potassium chloride	Samtec	ANVISA registration: 1559200010139	Parenteral drug
Procedure gloves	Descarpack	122401	Personal Protective Equipment (PPE)
Propofol (Provive)	União Química	ANVISA registration: 1049714490057	Anesthetic
Ringer lactate solution 500 mL bag	JP Farma	MS:1.0491.0061	Fluid therapy
Riohex 0.5% clorexidine alcohol solution	Rioquimica	218690356	Surgical asepsis
Riohex 2% clorexidine solution with surfactant	Rioquimica	218690356	Surgical asepsis
Scalp 21 G	Descarpack	421201	Surgical material
Shoe covers	generic	-	Personal Protective Equipment (PPE)
Sterile compresses	Cremer	ANVISA registration: 10071150065	Surgical material
Sterile gauze pad	Procitex	ANVISA registration: 80245210083	Surgical material
Sterile surgical drapes 140 cm x 90 cm	Venkuri	7010003	Surgical material
Sterile surgical drapes 150 cm x 190 cm	PolarFix	F00208	Surgical material
Sterile surgical gloves	Mucambo	CA: 39.317	Personal Protective Equipment (PPE)
Stethoscope	generic	-	Surgical equipment
Surgical cap	Descarpack	93201	Personal Protective Equipment (PPE)
Surgical gown	Descarpack	231101	Personal Protective Equipment (PPE)
Surgical mask	Descarpack	110701	Personal Protective Equipment (PPE)
Surgical scissors blunt-blunt	generic	-	Surgical instrument
Surgical scissors sharp-sharp	generic	-	Surgical instrument
Surgical stapler	Tradevet	-	Surgical material
Tekbond super glue	Tek Bond	78072720030	Surgical material
Thermometer	generic	-	Surgical equipment
Three Way Stopcock	Solidor	374	Surgical material
Tramadol hydrocloride	Agener União	MAPA registration: SP-000292-5.000002	Anesthetic
Transparent film dressing	Skinupper	ANVISA registration: 82307460014	Surgical material
Waterproof adhesive 10 cm x 4.5 cm	generic	364828	Surgical material