Video summary

Transferencia de embriones

Main summary

Key takeaways

Educational

Main ideas / lessons conveyed

  • The video is an instructional training (2nd edition) for embryo transfer (ET) and related lab steps, emphasizing that:

    • Embryo transfer is a commercial, established technique used for 30+ years.
    • It enables faster genetic improvement within cattle herds globally.
    • Success depends heavily on:
      • Technique
      • Correct timing
      • Correct placement
      • Careful handling (especially minimizing trauma to the uterus and protecting embryo viability)

  • The overall workflow taught spans:

    1. Donor preparation and embryo collection (including epidural anesthesia and day-7 horn flushing/horn washing)
    2. Embryo recovery and microscopic identification (staging/quality evaluation)
    3. Freezing/cryopreservation (ethylene glycol protocol, straw loading, controlled crystallization, storage)
    4. Transfer of frozen/thawed embryos to recipients (placement depth tied to corpus luteum and recipient cycle stage)
    5. Thawing considerations and immediate transfer practices

Methods and step-by-step instructions (detailed bullets)

A) Donor palpation and CL (corpus luteum) localization

  • Put on a plastic sleeve over the arm; remove fingertips; use tight-fitting latex gloves.
  • Palpate both ovaries of each donor to:
    • Identify the corpus luteum (CL)
    • Identify possible structures: CL, retained follicle, large cyst, or combinations
  • Mark the hip corresponding to the uterine horn associated with the CL:
    • If CL is right → mark right hip
  • Notes on ovary/CL identification:
    • ~65% of ovulations occur in the right ovary
    • Active ovary is often larger than the opposite ovary
  • Day-7 CL size reference:
    • Textbook CL: ~8 mm diameter, 3–5 mm high
    • Common larger CLs: 10–13 mm diameter, ~8 mm high

B) Prepare and use DVPS washing system (probe + filter + DVPS/double-phosphate saline buffer)

  • Select a 1 L DVPS bag (dual-phosphated saline buffer) containing surfactant polyvinyl alcohol AP (example: Biolift Advantage recovery medium).
  • Connect system:
    • Insert probe into wash bag; close clamps
    • Insert probe end into bag perforation
    • Connect filter to probe; attach flexible catheter
  • Purge and fill filter:
    • Press/turn to ensure proper insertion
    • Purge air and flow DVPS into filter until 20–30 mm of DVPS is in filter
  • Gravity wash method:
    • Lay bag horizontally; lower probe + filter below table surface
    • Let medium flow into filter container until 20–30 mm
    • Close clamps; return system to table surface

C) Prepare the double-lumen follicle catheter (for recovery)

  • Select and test a double-lumen follicle catheter
  • Example mentioned: Vortex catheter (long 23-inch silicone catheters emphasized for sterilization/reuse and designed for bovine retrieval)
  • Inflate balloon as part of setup:
    • Allow drops from probe into catheter
    • Insert sterile stainless steel stylet
    • Inflate balloon with 20 cc air, then remove 20 cc air (as described)
  • Completion steps:
    • Stretch catheter 0.5 inch toward the stylet connection port
    • Secure rear 0.25 inch end near stylet connection with hemostatic clamp

D) “Horn washing / horn washing technique” (day-7 embryos via horn-directed gravity flow)

Core concept

  • Use “horn washing” where recovery medium enters the uterus via gravity and is positioned to direct flow to where embryos are expected.

Key target

  • Day-7 embryos in superovulated donors are generally in the upper third of uterine horns.
  • Washing horns directs recovery medium toward the anterior half of horns.

D1) Epidural anesthesia preparation

  • Prepare epidural syringe:
    • Attach 18G x 1.5 inch needle to 6 cc syringe
    • Aspirate 5–6 cc of 2% lidocaine; cover needle
  • Pre-retrieval tray items:
    • Shoulder-length sleeves, latex gloves
    • Three hemostatic forceps
    • Lidocaine bottle, additional needles
    • Paper/pencil for recording structures

D2) Tail/infusion site cleaning and injection (as taught)

  • Remove feces/dirt from the tail where epidural applies.
  • Wash with liquid antiseptic (example: iodine solution), then rinse with clean water.
  • Positioning and injection:
    • Assume feeling with left hand (per explanation)
    • Put sleeve covering arm; remove manure from rectum
    • Grasp tail about 3 inches below rectum; lift slightly above horizontal
    • Use thumbnail of right ungloved hand to press tailbone region to locate a cleft between vertebrae (injection site)
    • Insert lidocaine needle deep into intervertebral space until bone contact
    • Inject lidocaine:
      • Back pressure early → withdraw needle slowly in ~1 mm intervals while continuing to press plunger
      • Correct dosing signs:
        • Tail remains flaccid after ~1 minute
        • Rectal peristalsis stops
  • (Some subtitle text is garbled; the main injection goal is correct epidural placement.)

E) Rectal preparation and uterine access

  • After epidural:
    • Remove soiled sleeve; put on clean fingerless sleeve + latex glove
    • Clean fecal matter from anal sphincter and vulva, rinse vulva after opening lips
  • Set bag height and lubricate:
    • Suspend 1 L DVPS bag with probe+filter so the bottom edge is about 1 meter above tail boom
    • Lubricate hand/arm with water (warning: other lubricants may harm embryos if they drip near catheter/transfer rod)
  • If rectum contains air:
    • Evacuate air by inserting fingers deeply until rectum closes in a tight ring
    • Grasp and gently pull back 2–3 times to expel air
  • Critical rule:
    • “All lubricants are toxic to the embryo.”

F) Locate cervix/horns and record ovarian/ovulation info

  • Identify cervix and uterine horns once inside donor.
  • Estimate:
    • number of OCL ovulations and retained follicles
    • ovarian volume (using length/width/height concept)
  • Use finger width to estimate volume quickly.
  • Record for later HFE dose adjustments (mentioned).

G) Catheter placement technique for horn washing

  • Remove catheter from packaging; open vulva; insert into vagina; pass through cervix “like AI.”
  • Uterine positioning:
    • Try to gather/pull back uterus toward pelvic floor
  • Placement immediately after catheter enters uterine body:
    • Position tract so that catheter is placed to bring right/left horn directly in front of catheter (not steering blindly)
  • Find palpable bifurcation:
    • Use middle finger downward pressure to identify the division point where horns separate
  • Advance:
    • Advance catheter tip 6–7 cm beyond bifurcation into target horn
  • Location and inflation determine success:
    • Location: balloon must seal against inner horn wall and not move during flushing
    • Inflation:
      • enough air for seal
      • too much can cause endometrial splitting
      • too little may cause balloon shift and poor recovery

G1) Balloon inflation protocol (assistant performs inflation)

  • Assistant inserts 20 cc syringe into catheter inflation valve with plunger at 20 cc mark
  • Press in 12 cc air
  • Operator palpates horn to locate partially inflated balloon:
    • If easy to identify → enough air
    • If not → add partial volumes of 1 cc until appropriate balloon feel/appearance
  • Typical amounts:
    • Cows: 14–16 cc
    • Virgin heifers: 12–13 cc
  • Safety threshold:
    • Placement should generally not need inflation > 16 cc (even in larger donors) to avoid complications

G2) After inflation: stylet removal and securing catheter

  • Remove hemostatic clamp holding catheter to stylet; attach clamp in front of inflation valve
  • Assistant disconnects air syringe from valve
  • Remove stylet:
    • Thumb+forefinger backstop to prevent catheter pulling back
    • Assistant gently twists and pulls stylet out
  • Prevent balloon detachment:
    • Use a second closed hemostatic clamp through ring on first clamp
    • Then clamp second onto donor hair near base of tail
  • Insert Y-probe foil adapter into catheter opening
  • Goal:
    • Fill/empty right horn several times, remove catheter, place back in left horn, repeat for remaining medium

H) Controlled filling/emptying and “milking” massage

  • Monitor the anterior half of horn (embryos at 7 days after estrus):
    • Place fingers around anterior half so upper small-diameter part rests in palm
  • During setup:
    • Close clamp leading to filter; open clamp allowing buffer entry
  • First fill:
    • Start conservative to establish flow and confirm fluid circulates into uterus and exits into filter
  • Monitor filling:
    • Gently tap upper third as it fills
    • When horn is full (tight/firm/swollen):
      • close inlet clamp
      • open outlet clamp to let liquid into filter
  • Massage (“milking”):
    • Immediately massage horn like “milking”
    • Start at smallest diameter area; move toward middle of horn
    • Repeat until no more liquid flows into filter
  • Timing guidance:
    • Fill: typically 60–90 seconds
    • Empty: 20–30 seconds
  • When 500 ml remains in hanging bag:
    • relocate catheter to other horn
  • If using certain filter types (e.g., zone filter):
    • transfer filter contents to a 90 mm round grid-bottom search dish

I) Filter rinsing / mesh search dish processing (recover embryos)

  • Rinse filter and mesh using a 20 cc two-piece plastic syringe with 16G needle
  • Filter rinsing volume:
    • 40–60 cc medium to properly rinse filter/mesh
  • Embryo safety restrictions:
    • Use only two-piece syringes if embryo-contact fluids are needed
    • Avoid syringes with butyl/black rubber stopper around plunger end (toxic surfactant coating can lower pregnancy rates)
  • Drain leaving ~6–7 mm liquid in filter
  • Spiral method:
    • Turn filter to form spiral in solution
    • Invert container; pour onto search plate
  • Wash filter walls:
    • Use syringe to wash walls; keep 6–7 mm liquid
  • Aspirate embryos from mesh:
    • Hold filter at eye level
    • Place syringe nipple into mesh
    • Slowly aspirate to draw mucus, debris, embryos
    • Expel into search plate
  • Bubble handling:
    • If bubbles exist in syringe, expel into an inverted cap for later search after bubbles dissolve
  • Microscopy search:
    • Plate on microscope stage
    • Embryos settle bottom in 1–2 minutes
    • Search magnification: 10–15x total
    • Move plate in a Z pattern starting upper-left grid
  • Removing mucus:
    • Use two 20G x 1.5 inch needles to immobilize and scrape mucus off embryo repeatedly

J) Embryo capture and staging/quality grading

  • When found:
    • Connect F and B catheter to a three-piece 1 cc syringe
    • Do not extract washing medium into syringe barrel
  • Prepare labeled dish(s):
    • Petrie dish lid/side labeled with donor name; multi-well plate example: six-well plate
  • Sorting approach:
    • Place embryos in first well; classify as search completes
    • Move “bad” embryos to later well; “good” embryos to another
    • Select for:
      • fresh transfer (“tea” mentioned—likely ET fresh)
      • freezing

K) Embryo transfer preparation (transfer medium + sterile filtration)

  • Withdraw 8–10 cc of transfer/maintenance medium into new 10–20 ml Cell Safe syringe
  • Optional but recommended:
    • sterile filter liquid embryos may contact using ET Media filter
  • After filtering:
    • discard first 4–5 drops
    • then add 6–8 drops to dish
  • Fill until ~65% full
  • Load embryo:
    • Use low-power microscopy
    • Aspirate a ~15 mm fluid column into catheter tip
    • Position catheter tip adjacent to embryo; aspirate slowly until embryo is 2–3 mm inside tip
    • Expel into dish gently:
      • expel liquid in catheter tip (exact units unclear due to subtitle errors, ~15–20 mentioned)
      • do not expel air into dish (prevents bubbles that embryos can adhere to and hinders development)

L) Embryo stage coding and quality assessment (freezing/transfer decisions)

  • Stage codes based on Appendix A (numerical codes 1–N)
  • Staging focuses on maturity of the main cell mass (blastomeres)
  • Common confusing stage guidance:
    • Stage 1 (non-fertile): single large cell enclosed in membrane
    • Stage 4 (morula): compact “bunch of grapes/soccer ball”
      • check edge border:
        • non-fertile border smooth/continuous
        • morula border wavy
  • Freezing-ready stages:
    • Stage 4 morula
    • Stage 5 early blastocyst
    • Stage 6 blastocyst
    • Stage 7 expanded blastocyst
    • Note: Stage 7 frozen has 5–10% lower pregnancy chance
  • Fresh-only caution:
    • Stage 3 early morula should not be frozen; can be transferred fresh on day 6
  • Quality scoring:
    • Based on debris/extruded blastomeres in perivitelline space (PV space) (between main mass and zona pellucida)
    • If no extruded blastomeres → quality level 1
    • If present → 2 or 3
  • Zona pellucida requirement for freezing:
    • Must be intact (no fractures/breaks)
  • Cryoprotectant (ethylene glycol, EG):
    • Necessary for survival through freezing
    • Limit exposure: longer than 10 minutes reduces survival
    • Last step is placing embryo in ethylene glycol

M) Freezing protocol overview (straw loading + controlled crystallization)

Preparation

  • Label straws; set freezer to -6°C
  • Prepare labeled petri dish/drops (letters EG for ethylene glycol dish)
  • Use syringe-filtered medium in receiving dish

Timeline (high level)

  • Timer set to 7 minutes (per protocol)
  • Minimize cryo exposure; follow crystallization timing windows

Per-embryo handling

  • Evaluate embryo stage/quality and record on C freezing certificate
  • Aspirate embryo into F and B catheter:
    • stop as soon as embryo enters catheter
  • Transfer into shaft plate; release into maintenance medium (minimal extra medium)
  • Start crystallization timing:
    • activate timer at start of cryo window
    • when 4 minutes remaining, load into 0.25 cc yellow straw

Straw loading technique (key mechanics)

  • Insert dry straw into three-piece syringe connector; create vacuum (art + practice)
  • Rinse straw first; avoid wetting connector initially
  • Create medium/air columns with embryo in final medium column:
    • column length ~45 mm
    • maintain ~45° straw angle to keep embryo centered
  • Create air gap:
    • yield a 6 mm air gap at end
  • Verify embryo position under microscope:
    • if embryo not centered within ~15 mm of the end farthest from cotton plug → reload
    • if centered, you may wet sealing plug

PVC saturation requirement (critical)

  • Ensure straw’s PVC powder is 100% saturated with shaft/medium:
    • pull plunger firmly and quickly once; stop when powder fully wets
  • Weak seal can burst during freezing

Seal and load into cryochamber

  • Insert labeled yellow sealing plug (centered; avoid bending/crushing)
  • Insert straw vertically into cryochamber slot(s)
  • Ensure freezer is -6°C
  • Wait until timer completes 7 minutes before crystallization

Crystallization procedure

  • Do not begin crystallization before the 7 minutes elapsed
  • Place clamp tips in liquid nitrogen bath for 20 seconds
  • Perform squeeze:
    • at ~1 mm below meniscus
    • for 10 seconds
  • Look for a frosted/white half-axis spot just below meniscus indicating correct protocol
  • Return straw to cryochamber
  • Timing rules:
    • Time between loading embryo on axis plate and starting crystallization must be ≤ 10 minutes
    • Load and crystallize one straw at a time (do not batch then crystallize together)

Storage

  • Freezing completes about 1 hour later, temperature ~-32°C
  • Transfer:
    • move straws from slots into goblet/cannula submerged in nitrogen
    • store cannula in nitrogen tank

N) Embryo transfer technique (frozen/thawed ET) — placement depth and recipient matching

  • Transfer success depends on:
    • Correct horn location relative to CL-associated horn
    • Correct deposit depth to minimize endometrial trauma
  • Average success target:
    • embryo deposited at least halfway into the CL-associated horn
    • “Halfway” = midpoint between palpable external bifurcation and end of horn
  • Better-than-average target:
    • place in upper third of horn with:
      • smallest diameter
      • furthest from cervix
      • least trauma
  • Avoid cervix damage:
    • allow cervix to shield sheath tip so it does not enter uterine wall
  • Horn deviation troubleshooting (left horn repeatedly misses):
    • If applicator/sheath deviates to right horn: place left palm against left horn side and push uterus to …

Original video