Protocol for an immunocompetent KPC mouse model of pancreatic cancer resection and recurrence for perioperative treatment studies

Summary

In mouse models of pancreatic ductal adenocarcinoma, surgical removal of orthotopic tumors typically involves removing the primary tumor along with the spleen. Here, we present a protocol for surgical removal of the tumor in the tail of the pancreas while preserving the spleen. We describe steps for implantation of mouse pancreatic cancer cells followed by the identification and removal of the orthotopic tumor. We also outline perioperative treatment strategies for tumor recurrence.

Graphical abstract

Highlights

• •Surgical steps for implanting KPC cells into the mouse pancreas
• •Surgical procedure for removal of KPC tumors from the mouse pancreas
• •Guidance on perioperative treatment strategies

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

In mouse models of pancreatic ductal adenocarcinoma, surgical removal of orthotopic tumors typically involves removing the primary tumor along with the spleen. Here, we present a protocol for surgical removal of the tumor in the tail of the pancreas while preserving the spleen. We describe steps for implantation of mouse pancreatic cancer cells followed by the identification and removal of the orthotopic tumor. We also outline perioperative treatment strategies for tumor recurrence.

Before you begin

This protocol outlines the establishment of an orthotopic pancreatic ductal adenocarcinoma (PDAC) model in immunocompetent mice for resection and recurrence. KPC1245 cells, derived from pancreatic tumors in LSL-KrasG12D; LSL-Trp53R172H; Pdx1-Cre mice are surgically implanted into the pancreas of C57BL/6 mice. Use newly thawed cells within a relatively low passage range and avoid prolonged in vitro culture, as changes in passage number may affect tumor growth and experimental reproducibility. Determine the number of mice required for each experimental group in advance, accounting for potential losses during tumor implantation, tumor establishment, resection, and postoperative recovery. If perioperative treatments are being tested, treatment schedules should be established before starting the experimental cohort. Ensure that all surgical rooms, instruments and supplies are available and in hand before starting. An approved standardized plan for perioperative monitoring, analgesia, recovery, and humane endpoints should also be in place and in accordance with the relevant institutional and regulatory guidelines for animal experimentation. In theory, this protocol could be adapted and optimized to other pancreatic cell lines and mice of different genetic backgrounds.

Innovation

Advanced metastatic disease is a main contributor to the poor clinical outcomes of pancreatic ductal adenocarcinoma (PDAC).¹ Surgical resection currently represents the only potentially curative treatment; however, only a small proportion of patients present with resectable disease.^2,3 Even after a successful surgery, patients can experience local or distant recurrence, suggesting that metastatic or residual tumor cells persist despite a complete resection.^4,5 As the use of neoadjuvant therapies increases and more patients become eligible for surgery, understanding the mechanisms driving postoperative recurrence has become increasingly important as multiple studies have demonstrated that the period following surgery is marked by a distinct state of immunosuppression, which cancer cells can exploit.^6,7,8 Preclinical mouse models have been essential for studying PDAC biology and testing therapeutic strategies. Many groups have developed orthotopic and surgically resectable PDAC models in immunocompetent mice to accurately reproduce the clinical scenario of tumor removal followed by disease recurrence. These models typically include a pancreatic transection and a partial or complete splenectomy.^9,10,11 However, preserving the spleen is particularly important for immunological studies as it maintains the immune competence of mice and avoids the physiological consequences associated with a splenectomy.^12,13 To address this limitation, this protocol describes a distal pancreatectomy with complete splenic preservation. This challenging dissection procedure protects the fragile splenic vessels while also achieving a complete removal of the pancreatic tumor. This innovation enables the use of fully immunocompetent mice, providing a physiologically relevant model to study immune responses to treatments in the perioperative time frame.

Institutional permissions

Approval from the University Animal Care Committee (UACC) or equivalent institutional approval is established before performing similar experiments. In this study, mouse experiments were carried out following the Canadian Council on Animal Care guidelines under the approved Université de Sherbrooke UACC protocol #2024-4708.

Preparation of cell culture media

Timing: 15 min

This step should be performed prior to KPC cell culture.

• 1.In a biological safety cabinet, prepare the cell culture media by adding 50 mL of Fetal Bovine Serum (FBS) and 5 mL of Penicillin-Streptomycin (P/S) to 445 mL of DMEM.
• 2.Warm the media to 37°C before use (approximately 1 h).

Note: Cell culture media can be stored at 4°C for 3–4 weeks.

Preparation of basement membrane matrix

Timing: 15 min

This step should be performed prior to orthotopic cell implantation.

CRITICAL: Avoid thawing on the bench at 20°C or in a 37°C water bath, because the basement membrane matrix starts to polymerize at around 10°C.

• 3.Thaw a 10 mL vial of basement membrane matrix at 4°C for a minimum of 4 h or place at 4°C the day before aliquoting.
• 4.
  Work on ice in a biological safety cabinet, prepare 20 aliquots of 500 μL of basement membrane matrix in Eppendorf tubes.

  • a.
    Thaw aliquots on ice before use (approximately 1 h).

Note: Basement membrane matrix aliquots can be stored at −20°C until the manufacturer expiry date.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Chemicals, peptides, and recombinant proteins
DMEM	Wisent Inc.	319-005-CL
Penicillin-Streptomycin, 100×	Wisent Inc.	450-201-EL
Fetal bovine serum	Wisent Inc.	090-150
Phosphate buffered saline (PBS)	Wisent Inc.	311-010-CL
Trypsin-EDTA 0.25%	Wisent Inc.	325-043-CL
Trypan Blue	Wisent Inc.	609-130-EL
Medical Oxygen	Linde Canada Inc.	OX M-K
Isoflurane	Fresenius Kabi	CP0406V2
Eye ointment	Optixcare	ORK001856
Buprenorphine HCL (SC) 0.6mg/mL	Chiron Compounding Pharmacy Inc.	N/A (prescription)
Petroleum jelly	Healthcare Plus	00741515
Basement membrane matrix (Matrigel)	Corning	354234
Ethanol	Greenfield Global Inc.	P025EAAN
Chlorhexidine Gluconate (Baxedin Pre-Op)	Omega Laboratories	L0000003
2% lidocaine HCl	Aspen Pharmacare Canada	02302438
0.5% bupivacaine	SteriMax Inc.	02443694
Saline	ICU Medical, Inc.	07983225
Critical commercial assays
Tumor dissociation kit, mouse	Miltenyi Biotec, Inc.	130-096-730
Experimental models: Cell lines
KPC1245 cell line	Gift from the laboratory of Dr. John Stagg, McGill University	KPC cells are also available for purchase from: https://cancertools.org/cell-lines/kpc-c57-bl6-genetic-background-153474/
Experimental models: Organisms/strains
C57BL/6J female mice, 6–8 weeks old	Jackson Laboratory	IMSR_JAX:000664
Other
GentleMACS dissociator, with Heaters	Miltenyi Biotec, Inc.	130-134-029
Hemacytometer	Hausser Scientific	3100
50 mL conical centrifuge tubes	Corning	352070
15 mL conical centrifuge tubes	Corning	352196
1.5 mL safe-lock tubes	Corning	MCT-150-C
Tissue culture dish, 100 × 20 mm	SARSTEDT	83.3902
Penlon Sigma Delta Vaporizer (anesthesia)	DISPOMED	990-VI5P-VISSCREW
Deltaphase Isothermal Pad	Braintree Scientific, Inc.	DP-PAD
Electronic Caliper	Mastercraft	58-6800-4
Ear punch	Fine Science Tools	24214-02
Spunlace Surgical Drapes	Jorgensen Laboratories, LLC	J0258
Mateo TL light microscope	Leica Microsystems	11526238
MedPro® Cotton Tipped Applicator, Sterile	AMG Medical	018-430
Insulin syringe, 0.5cc U100, 29g × 1/2″	ELIMEDICAL	05V2913VB
Surgical scissors	Almedic	M04-1240
Surgical forceps (2)	Fine Science Tools	11051-10
Crile Hemostatic Forceps - 5-1/2″, Straight	Sklar	17-3055
Stainless steel scalpel handle (#3)	Fisherbrand	12-000-163
4-0 absorbable suture	Ethicon	VCP442H
Sterile gauze pad	Fisherbrand	22-028-556
Surgical Cautery Accu-Temp® Fine Tip High Temperature	Beaver-Visitec International	8442000
PIPETMAN P200	Gilson Inc.	F144058M
200 μL Graduated Filter Tips, Sterile	UltiDent Scientific	50-6013
9mm stainless wound clips (AutoClip®)	Fine Science Tools	12022-19
9mm wound stapler (AutoClip® Applicator)	Fine Science Tools	12020-09
Staple remover (AutoClip® Remover)	Fine Science Tools	12023-00

Materials and equipment

Cell culture media

Reagent	Final concentration	Amount
DMEM	N/A	445 mL
Fetal Bovine Serum (FBS)	10 %	50 mL
Penicillin-Streptomycin (P/S)	1 %	5 mL
Total	N/A	500 mL

Store at 4°C for 3–4 weeks.

KPC injection mix

Reagent	Final concentration	Amount
Basement membrane matrix	50 % (v/v)	10 μL
KPC cells at 100000 cells/mL	50 % (v/v)	10 μL
Total	50000 cells/mL	20 μL

Store on ice or at 4°C for a maximum of 4 h.

Step-by-step method details

Culture KPC cells for 2–3 passages

Timing: 1 h every 2 days

The following steps describe the culture of KPC1245 cells.

CRITICAL: These cells grow rapidly and typically become confluent in 2 days. Letting the cells grow beyond 90% confluency will reduce the effectiveness of this protocol.

CRITICAL: Use newly thawed cells within a relatively low passage range and avoid prolonged in vitro culture as changes in passage number may affect tumor growth and experimental reproducibility.

Note: The growth characteristics described here are specific to KPC1245 and may not be representative of all KPC-derived cell lines. Tumors developed from these cells have been well-characterized and are typically used to study therapeutic responses to immunotherapies and chemotherapies.^14,15,16

Note: The following steps should be performed using aseptic techniques in a biosafety cabinet.

• 1.Prewarm sterile PBS and cell culture media at 37°C.
• 2.
  Thaw KPC cells from a cryovial (approximately 1 million cells) into a cell culture dish.

  • a.
    Fill a 15 mL falcon with 9 mL of cell culture media.
  • b.
    Thaw the cryovial of cells in a 37°C water bath (approximately 30 to 60 s).

    • i.
      When the cryovial has thawed, but still cold, gently resuspend the cells in the prepared 15 mL conical tube.
  • c.
    Centrifuge content at 350 × g for 5 min.

    • i.
      Decant the supernatant without disturbing the cell pellet.
    • ii.
      Resuspend the cells in 10 mL of cell culture media.
  • d.
    Gently transfer this new 10 mL of KPC cell suspension to a 100 mm cell culture treated dish.

    • i.
      Place the cell dish in a humidified cell incubator at 37°C.

Note: The confluency of KPC cells will reach 70%–90% confluency every 2 days and should be passaged before they become more than 90%–100% confluent.

• 3.
  Subculture the cells following routine cell culture procedures.

  • a.
    Remove the cell culture media from the cell culture dish.
  • b.
    Wash the cells with 10 mL of warm PBS for 30 s.

    • i.
      Remove the PBS from the cell culture dish.
  • c.
    Add 1 mL of 0.25% trypsin-EDTA to the cell culture dish.

    • i.
      Place the dish in the cell incubator at 37°C for 2 min.
  • d.
    Observe the KPC cells under a microscope that the cells are beginning to detach from the cell culture dish.
  • e.
    Inhibit the trypsin reaction with 10 mL of cell culture media.

    • i.
      Quickly rinse the dish several times with the media.
    • ii.
      Collect the KPC cell suspension in a 15 mL conical tube.
  • f.
    Centrifuge content at 350 × g for 5 min.

    • i.
      Decant the supernatant without disturbing the cell pellet.
    • ii.
      Resuspend the cells in 10 mL of cell culture media.
  • g.
    To subculture the KPC cells, replate 1 mL of the cell suspension volume in a new cell culture dish containing 9 mL of cell culture media.

    • i.
      Place the dish in the 37°C cell incubator.
    • ii.
      If you are implanting on the same day, place the cells on ice and proceed to the next step.

Preparation of the KPC injection mix

Timing: 1 h

The following steps describe the preparation of KPC cells for injection in mice.

CRITICAL: Always work on ice to prevent the polymerization of the basement membrane matrix.

• 4.
  Prepare 10 mL of KPC cells at a concentration of 100 000 cells/mL.

  • a.
    Combine 900 μL of PBS and 100 μL of cells prepared at step 3 in an Eppendorf tube.
  • b.
    Count cell dilution with a classic hematocytometer.

    • i.
      Withdraw 10 μL of the cell suspension and combine 10 μL of Trypan blue in another Eppendorf tube.
    • ii.
      Count the live cells following standard cell counting protocols.
    • iii.
      Multiply the result by the dilution factor of 20 using the formula below.

      $$Totalcells/mL=\frac{Totalcellscounted\ast dilutionfactor\ast 10000}{\#ofsquarescounted}$$
  • c.
    Resuspend the cells in cell culture media at 100 000 cells/mL.

    • i.
      Place the cells on ice.

      Note: Determine the resuspension volume using the equation C₁V₁ = C₂V₂. In this formula, C₁ represents the measured cell concentration calculated at step 4, V₁ is the volume required of the cell suspension, C₂ is the desired final cell concentration (100 000 cells/mL), and V₂ is the final total volume of the suspension (10 mL). For instance, if the measured cell concentration is 2 million cells/ml, 500 μL of cells should be resuspended in 9.5 mL of cell culture medium for a total final volume of 10 mL.

• 5.
  Prepare the KPC injection mix at a ratio of 1:1 with basement membrane matrix for a final concentration of 50 000 cells/mL.

  Note: The injection volume is 20 μL per mouse (1000 KPC cells per mouse).

  • a.
    Thaw the required number of already prepared 500 μL aliquots of basement membrane matrix on ice before use (approximately 1 h).

    • i.
      Place basement membrane matrix on ice.

      Alternatives: Other animal-derived basement membrane extracts, such as Cultrex (Bio-Techne), could theoretically be used as substitutes for Matrigel but have not been validated in this model.

  • b.
    Working on ice, prepare the required volume of injection mix by gently mixing.
  • c.
    Gently mix an equal volume of cells prepared at step 4 (100 000 cells/mL) with an equal volume of basement membrane matrix.

    • i.
      Place the injection mix on ice.

      Note: Determine the volume of injection mix by multiplying the number of mice to inject by 20 (20 μL is the injection volume). That number is then multiplied by 3 to account for pipetting error, premature gelation, and unsuccessful injections. For example, if the plan is to inject 20 mice, 600 μL of cells should be resuspended with 600 μL of basement membrane matrix for a total final volume of 1200 μL.

Orthotopic implantation of KPC cells

Timing: 1 day

These steps outline how to surgically implant KPC cells into the pancreas of a C57BL/6 mouse using a small incision beneath the left rib cage. It is important to adhere to the surgical guidelines approved in your institution’s UACC protocol.

Note: Each procedure initially takes around 15–25 min to complete alone, all the way down to under 10 min after gaining experience.

Note: Before starting the procedure, ensure that the anesthesia system and the isoflurane scavenging unit are functioning properly. Throughout the surgery, carefully monitor the mouse for signs of excessive or insufficient anesthesia and adjust the isoflurane flow rate as needed.

Note: The mouse should remain on a heating pad during the surgery and throughout the recovery period.

CRITICAL: All surgical instruments must be thoroughly cleaned and sterilized before use.

CRITICAL: Keep injection mix on ice to prevent the polymerization of the basement membrane matrix.

• 6.
  Anesthetize the mouse before surgery.

  • a.
    Turn on the oxygen flow rate to 1–2 L/minute and the isoflurane flow rate to 2–3% for the induction chamber.

    • i.
      Place a mouse in the anesthesia induction chamber.
  • b.
    Monitor the mouse’s respiratory rate and appearance until anesthetized per pre-established UACC guidelines.

    • i.
      Turn down the flow rate of the isoflurane to 1–2%.
    • ii.
      Remove the mouse from the induction chamber.
  • c.
    Place the head of the mouse in the anesthesia cone situated on a heated pad.

    • i.
      Perform a toe pinch to confirm that the mouse is adequately sedated.

Note: If the mouse is adequately sedated, there would be no movement generated by the pinch. If the pinch triggers a reflex, wait a couple of s before proceeding and check again.

• 7.
  Prepare the mouse for surgery.

  • a.
    Identify the mouse using an ear punch per UACC guidelines.
  • b.
    Gently apply ointment to both eyes.
  • c.
    Inject long-acting (SR) buprenorphine subcutaneously at a dose of 1 mg/kg.

    Note: Under UACC guidelines, long-acting (SR) buprenorphine must be stored in a securely locked cabinet and administered only by trained veterinarians or technicians. Consult UACC guidelines for proper dosage and usage of the drug.

  • d.
    Position the mouse so that it is lying on its right side on the warm heating pad.

    • i.
      Tape the mouse at the shoulders and legs in a slightly stretched position.
  • e.
    Apply chlorhexidine to clean the incision site.
  • f.
    Apply 70% ethanol to clean the incision site.
  • g.
    Apply a small amount of sterile petroleum jelly with a cotton swab at the incision site.

    • i.
      Push aside the hair to facilitate the surgery.
  • h.
    At the incision site, inject 2% lidocaine HCl at a dose of 7 mg/kg and 0.5% bupivacaine at a dose of 3.5 mg/kg.

    Note: Determine the volume of drugs needed using the equation C₁V₁ = C₂V₂. This example assumes a mouse weight of 19 g, which is equivalent to 0.019 kg, an injection volume of 0.1 mL, and an experimental plan to inject 20 mice. In this formula, C₁ represents the 2% lidocaine HCl which is equivalent to 20 mg/mL, V₁ is the volume of lidocaine required for the preparation, C₂ is the desired final drug concentration (7 mg/kg × 0.019 kg ÷ 0.1 mL = 1.33 mg/mL), and V₂ is the final volume of the drug preparation (20 mice x 0.1 mL per injection = 2 mL). For the other drug, C₁ represents the 0.5% bupivacaine which is equivalent to 5 mg/mL, V₁ is the volume of bupivacaine required for the preparation, C₂ is the desired final drug concentration (3.5 mg/kg × 0.019 kg ÷ 0.1 mL = 0.665 mg/mL), and V₂ is the final volume of the drug preparation (20 mice × 0.1 mL per injection = 2 mL). In this example, 133 μL of 2% lidocaine HCl and 266 μL of 0.5% bupivacaine are resuspended with 1601 μL of saline for a total final volume of 2 mL. Consult UACC guidelines for proper dosage and usage of the drugs.

• 8.
  Perform the mouse surgery (video tutorial in Methods video S1).

  • a.
    Make a 1.5 cm incision in the skin at the level of the third left mammary gland (left upper abdomen) (Figure 1A).
  • b.
    Using surgical forceps, elevate the muscle fascia away from the peritoneal contents.

    Note: The dark red color of the spleen can be easily located through the semi-transparent peritoneum. This provides an approximate location for the incision site through the peritoneum.

  • c.
    Make a 1 cm transverse opening in the peritoneum with surgical scissors.

    • i.
      Place one 4–0 absorbable suture through the peritoneum of the mouse to hold the abdominal cavity open.
    • ii.
      Locate the pancreas and identify its tail on the medial side of the spleen.
  • d.
    Gently exteriorize the pancreas by pulling it out by the tail using forceps.

    • i.
      Gently pull and spread the pancreas, so it sits extracorporeal on a stainless-steel scalpel handle sitting on the abdomen of the mouse (Figure 1B).

      CRITICAL: In case of light pancreatic or splenic bleeding, use a sterile cotton swab to apply a gentle pressure on the bleeding vessel for 10–30 s. Continue with the procedure once bleeding stops.

      CRITICAL: In case of heavy pancreatic or splenic bleeding, apply a firm and sustained pressure for 30–60 s with a sterile gauze pad or a sterile cotton swab. Once the bleeding slows, try to visualize the vessel. If the bleeding has almost stopped, continue applying pressure for 30–60 s. If the bleeding has not stopped, use an electrocautery to carefully cauterize the bleeding vessel. Because mice have very low blood volume, if symptoms like pale organs or labored breathing appear, it is more humane to terminate the procedure according to established UACC protocol.
• 9.
  Implant KPC cells in the pancreas (video tutorial in Methods video S1).

  • a.
    Gently mix the injection mix using a P200 pipette.

    • i.
      Aspirate 20 μL using a sterile single-use insulin 0.5 mL syringe.
  • b.
    Carefully implant the cells in the tail of the pancreas by injecting 2–3 mm deep towards the head of the pancreas (Figure 1C).

    • i.
      Wait 1 second and remove the needle slowly.
    • ii.
      Inspect the injection site to ensure there is very little to no leakage of cells.

      CRITICAL: Ensure that the needle does not completely pierce the pancreas before injecting the cells. A basement membrane matrix bubble should form instantly and a visual confirmation of the bubble results in a successful tumor engraftment (video tutorial in Methods video S1).

      CRITICAL: In case of a minor or complete cell leakage where the needle completely transpierced the pancreas, soak the spilled cells with a cotton swab to prevent cells from entering the abdominal cavity. Wash the scalpel with 100 μL of PBS, dry with a cotton swab, and attempt another injection, ensuring not to inject directly in the same location.

  • c.
    Gently lift the stainless-steel scalpel handle to allow the pancreas to gently slide back into the peritoneal cavity.

    Note: If the pancreas is stuck to the stainless-steel scalpel handle, give it a gentle push with the tip of your forceps.

  • d.
    Inspect the pancreas to ensure it has been repositioned in the abdominal cavity.
• 10.
  Prepare to close the peritoneum and skin with the initiated stitch of 4–0 absorbable suture (video tutorial in Methods video S1).

  • a.
    Tie 3 knots and cut the tails short (Figure 1D).
  • b.
    Remove the tape holding the mouse.
  • c.
    Close the skin using 2–3 sterile stainless wound clips (9mm) (Figure 1E).
  • d.
    Apply chlorhexidine to sterilize the surgery site.
  • e.
    Turn the mouse around.

    • i.
      Administer a subcutaneous injection of 500 μL of saline solution.
  • f.
    Remove the mouse from the anesthesia cone.

    • i.
      Place the mouse in a recovery cage positioned on a heated pad.
    • ii.
      Observe the animal during the recovery following UACC guidelines.
  • g.
    Clean surgical tools for 30 s in 70% ethanol.
  • h.
    Rinse surgical tools for 30 s in sterile PBS.
  • i.
    Proceed to the next mouse.

Figure 1.

Orthotopic implantation of KPC cells in mice pancreas

(A) Area of incision in the left abdomen.

(B) Exteriorization of the pancreas with forceps.

(C) Implantation of KPC cells in the tail of the pancreas.

(D) Closing of the peritoneum with sutures.

(E) Closing of the skin with stainless wound clips.

(F) Expected tumor volumes at post-implantation days indicated, data are shown as mean ± SD of eight biological replicates (n = 8).

Monitoring of KPC tumor growth

Timing: 1 h every 2 days

These steps outline how the mice should be monitored as per UACC protocol after the tumor implantation.

Note: The KPC orthotopic mouse model is a very aggressive model that requires follow-up assessments of mice well-being at least every two days. As the condition progresses, daily monitoring may also become necessary.

• 11.
  Follow-up on mice’s well-being every two days.

  • a.
    Evaluate mice according to a stress score (Figure S1).
  • b.
    Weigh mice and examine their general well-being.

Note: Any animal that loses more than 20% of its initial body weight should also be humanely euthanized in accordance with the UACC guidelines.

• 12.
  Monitor tumors using the palpation method.

  • a.
    Restrain the mouse gently to expose the abdominal area.

    • i.
      Palpate the abdomen gently with the tip of your fingertips.
    • ii.
      Assess for the presence of discrete firm structures.
  • b.
    Record the tumor measurement (Figure 1F).

    • i.
      Gently pinch up the tumor.
    • ii.
      Measure its long and short sides with calipers through the skin.
    • iii.
      Obtain an estimate of the tumor volume in mm³ using the formula below.

      $$V=0.5\ast longside\ast {shortside}^2$$

      CRITICAL: The palpation method provides only an approximate assessment of tumor size and should not be used for quantitative tumor measurements intended for publication. Even when performed correctly, measurements may vary by 2–3 mm per side due to the thickness of the mouse’s skin.

      Note: If direct visualization and measurement of pancreatic tumors is required, perform exploratory laparotomy no more than once a week in accordance to UACC guidelines. Since this is an invasive procedure that may add to animal stress and discomfort, especially during a treatment cohort, the frequency should be minimized and limited to what is relevant as per UACC guidelines.

      Optional: Perform the exploratory laparotomy (Figure 2).

  • c.
    Re-open mouse skin and peritoneum as described in steps 6–8.
  • d.
    Gently exteriorize the pancreas by pulling it out by the tail using forceps.
  • e.
    Gently position the sterile caliper jaws against opposite margins of the tumor.

    • i.
      Measure its long and short sides with calipers.
    • ii.
      Obtain an estimate of the tumor volume in mm³ using the formula below.

      $$V=0.5\ast longside\ast {shortside}^2$$

      CRITICAL: Avoid direct compression or excessive touching of the pancreas with the caliper, as squishing the pancreas may cause tissue damage and compromise the experiment.

  • f.
    Close the peritoneum and skin as described in step 10.
  • g.
    Sterilize the caliper for 30 s in 70% ethanol.
  • h.
    Rinse the caliper for 30 s in sterile PBS.

    • i.
      Proceed to the next mouse.

      Note: Where available, tumors can also be monitored using small-animal computed tomography (CT) to enable non-invasive visualization and measurement of tumor size.

Figure 2.

Exploratory laparotomy performed on day 6 post-implantation

Neoadjuvant therapy window of treatment (optional)

Timing: 1 h for up to 3 days

These steps outline the timing to administer neoadjuvant treatments as this period is suitable for modeling clinically relevant interventions prior to surgery. It is important to adhere to approved UACC protocols prior to administration of treatments.

Note: Depending on the experimental design, cohorts may receive no treatment, systemic chemotherapy, or immunotherapy during the neoadjuvant window. Alternative multi-agent chemotherapy regimens, such as FOLFIRINOX can also be evaluated; however, careful monitoring is required due to potential toxicity.

• 13.Allow tumors to establish for 3 days post-implantation.

Note: This time point corresponds to a localized well-established tumor.

• 14.
  Assess mice prior to treatments.

  • a.
    Assess overall health as described in step 11.
  • b.
    Confirm tumor engraftment through abdominal palpation as described in step 12.
• 15.
  Administer the neoadjuvant therapy.

  • a.
    Randomize mice into desired experimental groups.
  • b.
    Continue treatment over the defined neoadjuvant window (day 3 to day 5) (Figure 3A).

CRITICAL: Treatments should not be given past day 5 post-implantation to prevent adverse effects and toxicity before the resection surgery.

Note: Intraperitoneal (IP) administration of FOLFIRINOX chemotherapy was well tolerated and safe in this resection model (Figure 3B).

• 16.Monitor mice during treatment as described in step 11–12.

CRITICAL: Intensive chemotherapy regimens may result in significant toxicity and early mortality in mice cohorts. One or two doses of neoadjuvant chemotherapy are sufficient when studying mechanisms in the perioperative period.

Figure 3.

Perioperative treatment windows

(A) Schematic of a potential perioperative treatment plan.

(B) Kaplan-Meier survival curve of mice having surgery only compared to surgery and one neoadjuvant intraperitoneal dose of FOLFIRINOX (Ca²⁺ folinate 50 mg/kg, oxaliplatin 2.5 mg/kg, irinotecan 25 mg/kg, 5-fluorouracil 25 mg/kg) on day 5 post-implantation (n = 12), ∗∗p < 0.01, logrank test.

Resection of distal pancreatic KPC tumors

Timing: 1 day

These steps outline how to perform a distal pancreatectomy with a complete splenic preservation of the pancreatic KPC tumor using a small incision beneath the left rib cage of the mouse around day 8 post-implantation. It is important to adhere to the surgical guidelines approved in your institution’s UACC protocol.

Note: Each procedure initially takes around 15–25 min to complete alone, all the way down to under 10 min after gaining experience.

Note: Before starting the procedure, ensure that the anesthesia system and the isoflurane scavenging unit are functioning properly. Throughout the surgery, carefully monitor the mouse for signs of excessive or insufficient anesthesia and adjust the isoflurane flow rate as needed.

Note: The mouse should remain on a heating pad during the surgery and throughout the recovery period.

Note: The pancreas should contain an easily identifiable primary tumor, while adjacent local regions in the abdomen should be free of invasion.

CRITICAL: All surgical instruments must be thoroughly cleaned and sterilized before use.

CRITICAL: Ensure consistent surgical techniques across all groups to minimize variability.

• 17.
  Anesthetize the tumor-bearing mouse.

  • a.
    Turn on the oxygen flow rate to 1–2 L/minute and the isoflurane flow rate to 2–3% for the induction chamber.

    • i.
      Place a mouse in the anesthesia induction chamber.
  • b.
    Monitor the mouse’s respiratory rate and appearance until anesthetized per pre-established UACC guidelines.

    • i.
      Turn down the flow rate of the isoflurane to 1–2%.
    • ii.
      Remove the mouse from the induction chamber.
  • c.
    Place the head of the mouse in the anesthesia cone situated on a heated pad.

    • i.
      Perform a toe pinch to confirm that the mouse is adequately sedated.

Note: If the mouse is adequately sedated, there would be no movement generated by the pinch. If the pinch triggers a reflex, wait a couple of s before proceeding and check again.

• 18.
  Prepare the tumor-bearing mouse for surgery.

  • a.
    Gently apply ointment to both eyes.
  • b.
    Inject long-acting (SR) buprenorphine subcutaneously at a dose of 1 mg/kg.

    Note: Under UACC guidelines, long-acting (SR) buprenorphine must be stored in a securely locked cabinet and administered only by trained veterinarians or technicians. Consult UACC guidelines for proper dosage and usage of the drug.

  • c.
    Position the mouse so that it is lying on its right side on the warm heating pad.

    • i.
      Tape the mouse at the shoulders and legs in a slightly stretched position.
  • d.
    Remove stainless wound clips (9mm) with a staple remover.
  • e.
    Apply chlorhexidine to clean the incision site.
  • f.
    Apply 70% ethanol to clean the incision site.
  • g.
    Apply a small amount of sterile petroleum jelly with a cotton swab at the incision site.

    • i.
      Push aside the hair to facilitate the surgery.
  • h.
    At the incision site, inject 2% lidocaine HCl at a dose of 7 mg/kg and 0.5% bupivacaine at a dose of 3.5 mg/kg.

    Note: Determine the volume of drugs needed as outlined in step 7. Consult UACC guidelines for proper dosage and usage of the drug.

• 19.
  Perform the surgical resection of tumor-bearing mouse (video tutorial in Methods video S2).

  • a.
    Make a 1.5 cm incision in the skin at the same area where mice were originally implanted with KPC cells (Figure 4A).
  • b.
    Using surgical forceps, elevate the muscle fascia where stitches where sutures closed the peritoneal wound.
  • c.
    Make a 1 cm transverse opening in the peritoneum with surgical scissors.

    • i.
      Place one 4–0 absorbable suture through the peritoneum of the mouse to hold the abdominal cavity open.
    • ii.
      Find the tumor-bearing pancreas and gently pull it out by the tail using forceps, so it sits extracorporeal on a sterile gauze pad (Figure 4B).

      Note: The use of cotton gauze pads allows the pancreas to stick to a surface which allows a greater finesse for the following steps.

      CRITICAL: In case of light bleeding, use a sterile cotton swab to apply a gentle pressure on the bleeding vessel for 10–30 s.

      CRITICAL: In case of heavy bleeding, apply a firm and sustained pressure for 30–60 s with a sterile gauze pad or a sterile cotton swab. Once the bleeding seems to slow down, try to visualize the vessel. If the bleeding has almost stopped, continue applying pressure for 30–60 s. If the bleeding has not stopped, use an electrocautery to carefully cauterize the bleeding vessel. Because mice have very low blood volume, if symptoms like pale organs or labored breathing appear, it is more humane to terminate the procedure according to established UACC protocol.

  • d.
    Clamp down the end of the tumor using a hemostat.

    • i.
      Stretch out the tumor-bearing pancreas.
    • ii.
      Visualize the healthy tissue and the tumor margin.
  • e.
    Gently pass a thread of 5–0 absorbable sutures under the tumor until it reaches the healthy margin of the pancreas.

    • i.
      Go 5 mm past the tumor in the healthy pancreatic margin (Figure 4C).

      CRITICAL: Ensure to preserve the pancreatic and splenic vasculature while ensuring complete tumor separation before doing the ligature (see Methods video S2).

  • f.
    Tightly ligature the tumor with 4 knots and cut the tails short.
  • g.
    Using surgical scissors, remove the tumor in one or two gentle cuts distal to the ligature (approximately in the middle of the 5mm healthy margin) (Figure 4D).

    Note: During this step, the resected tumor can be immediately collected and placed in PBS. Due to the highly enzymatic nature of pancreatic tissue, single-cell dissociation within 2 h of tumor resection is critical to preserve cell viability and integrity for downstream immune profiling applications (flow cytometry and transcriptomic analysis). The use of a commercial dissociator and kit (GentleMACS, Miltenyi Biotec) facilitates rapid and standardized tissue dissociation, helping to maximize cell yield and integrity for downstream analyses.

  • h.
    Cauterize the cut using a high-temperature cauterization tool (Figure 4E).

    CRITICAL: Ensure that the high-temperature cauterization tool does not go too close to the ligation as it could burn the thread.

    CRITICAL: This step is important to minimize bleeding and reduce the risk of pancreatic enzyme leakage in the abdomen.

  • i.
    Using surgical forceps, gently return the pancreas into the peritoneal cavity.

    • i.
      Inspect the pancreas to ensure it has been repositioned in the abdominal cavity.
    • ii.
      Inspect for bleeding.
• 20.
  Prepare to close the peritoneum and skin with the initiated stitch of 4–0 absorbable suture.

  • a.
    Tie 3 knots and cut the tails short.
  • b.
    Remove the tape holding the mouse.
  • c.
    Close the skin using 2–3 sterile stainless wound clips (9mm).
  • d.
    Apply chlorhexidine to sterilize the surgery site.
  • e.
    Turn the mouse around.

    • i.
      Administer a subcutaneous injection of 500 μL of saline solution.
  • f.
    Remove the mouse from the anesthesia cone.

    • i.
      Place the mouse in a recovery cage positioned on a heated pad.
    • ii.
      Observe the animal during the recovery following UACC guidelines.
  • g.
    Clean surgical tools for 30 s in 70% ethanol.
  • h.
    Rinse surgical tools for 30 s in sterile PBS.

    • i.
      Proceed to the next mouse.

Figure 4.

Resection of distal pancreatic KPC tumors

(A) Area of incision in the left abdomen.

(B) Clamping down the end of the tumor using a hemostat.

(C) Passing sutures under the tumor and ligature in the healthy margin of the pancreas.

(D) Cutting the tumor.

(E) Cauterizing the tumor.

Adjuvant therapy window of treatment (optional)

Timing: 1 h for up to 3 days

These steps outline the timing to administer adjuvant treatments as this period is suitable for modeling clinically relevant interventions post-resection surgery. It is important to adhere to approved UACC protocols prior to administration of treatments.

Note: After resection, mice may be observed, continue receiving a similar adjuvant immunotherapy, or be assigned to a new adjuvant treatment, depending on the study's design.

CRITICAL: In this model’s very early adjuvant treatment window, additional chemotherapy treatment regimens may become too toxic for mice.

• 21.Allow mice to recover for at least 3 days after the surgical resection.
• 22.Assess mice prior to treatments as described in step 11.
• 23.
  Administer the adjuvant therapy.

  • a.
    Continue treatment over the defined adjuvant window (day 11 to day 13) (Figure 3A).
  • b.
    Monitor mice during treatment as described in step 11–12.

Monitoring for tumor recurrence

Timing: 1 h every 2 days

These steps outline how mice should be monitored as per UACC protocol after the surgical resection of the primary pancreatic tumor.

Note: The recurrence phase theoretically begins on the day of the tumor resection and since the KPC orthotopic mouse model is a very aggressive model, it requires follow-up assessments of mice well-being at least every two days.

Note: Without treatment, tumor recurrence in the abdomen should be distinguishable through palpation of the abdomen 5–10 days after the resection surgery.

• 24.Follow-up on mice well-being every two days as described in steps 11–12.

Optional: Visualize the abdominal cavity and assess the pancreas and surrounding tissues for evidence of tumor recurrence and metastasis as described in step 13.

• 25.
  Define the experimental endpoint.

  • a.
    Record overall survival as the primary endpoint for evaluating the effects of the administered therapies.
  • b.
    Select additional endpoints to address specific mechanistic objectives of the study.

Expected outcomes

Mice implanted orthotopically with KPC cells using this protocol are expected to develop pancreatic tumors of approximately 300 mm³ within 8 days. At this stage, distal pancreatectomy can be performed, providing a model to investigate tumor recurrence following primary tumor removal. The disease progression without including the resection surgery can result in significant morbidity as early as day 14 post-implantation.

After the resection surgery, mice develop detectable primary tumor recurrence within 5–10 days, with some exhibiting peritoneal metastasis and/or ascites at later stages of tumor progression. In this model, neoadjuvant chemotherapy administered prior to resection has shown to improve survival outcomes (Figure 3A), which recapitulates aspects of current clinical outcomes.^17,18

This protocol can be used by experimenters to evaluate the tolerability and potential effects of different combination treatment regimens on tumor recurrence and survival. Perioperative treatment strategies are outlined to evaluate therapeutic approaches aimed at preventing or delaying tumor recurrence (Figure 3B).

Limitations

Cell and tumor growth may vary under both in vitro and in vivo conditions. Very rare instances of metastasis and local invasion to the spleen and abdominal wall have been documented prior to the resection. Mice exhibiting evidence of metastatic disease prior to the planned resection should be excluded from subsequent recurrence analyses. Additionally, variability in tumor establishment can influence how tumors respond to treatment, and treatment-related toxicity may confound survival outcomes.

Troubleshooting

Problem 1

Air bubbles form in the basement membrane matrix while preparing aliquots or the injection mix (step 5).

Potential solution

To avoid bubbles, handle the viscous solution with slow, gentle pipetting, taking care not to introduce air from the pipette tip into the gel mixture. If too many bubbles are introduced, rest on ice for 15 min to allow bubbles to rise to the surface of the solution.

Problem 2

The basement membrane matrix solution or injection mix begins to polymerize too early (step 5).

Potential solution

To avoid this issue, keep the basement membrane matrix solution on ice at all times, including during pipetting and dispensing. If the stock solution polymerizes, it must be remade. If polymerization occurs during cell preparation or surgery, discard the affected gel and prepare a new mix, ensuring the solution remains on ice throughout. A great way to avoid this issue is to prepare several aliquots of injection mix instead of one big volume solution (5 x 1 mL aliquots instead of 1 x 5 mL aliquots).

Problem 3

Cells do not become confluent after 2–3 days (Step 1).

Potential solution

This can happen for several reasons. Cells that become overconfluent may enter cell cycle arrest and fail to proliferate effectively. In such cases, thaw a new vial of cells and restart the culture protocol. Also, ensure that the incubator is properly humidified and that there is no evidence of contamination. It is not recommended to implant cells immediately after they thaw. Instead, allow them to recover and reach the exponential growth phase for at least 2–3 passages before implantation.

Problem 4

Tumor sizes observed in vivo show considerable variation (Step 12).

Potential solution

Although some variation in tumor size is normal, excessive differences can be problematic. Prior to injecting KPC cells in the pancreas, thoroughly mix the cells by pipetting to ensure even distribution. It is important to note that different clones of the KPC cell lines may exhibit differences in growth characteristics and tumor-forming capacity. Therefore, the specific KPC cell line and its source should be considered when interpreting tumor growth variability. In extreme cases, an in vivo growth curve kinetic experiment (Figure 1F) may be necessary to account for the variability of newly purchased cells. Contact L.-H.T. for more details.

Problem 5

Excessive bleeding during implantation or during resection surgery (Step 8 and Step 20).

Potential solution

Rough handling of the pancreas and surrounding tissues and prolonged surgical time may increase the risk of excessive bleeding due to poor hemostasis. Using fine surgical instruments and minimal manipulation of the organs greatly reduces the risk of bleeding. Using gentle pressure or a cautery tool remains the best option to control the bleeding; additional absorbable hemostats like SURGICEL can also be used. Practicing sham surgeries before experimental cohorts should improve speed and surgical skills.

Problem 6

High mortality rates after the resection surgery (Step 20).

Potential solution

This can happen for several reasons. Inadequate analgesia or overdosing, hypothermia during or after surgery, fluid loss or pancreatic enzyme leakage. Ensure optimized analgesic regimens, maintain a proper heat source during the surgery, administer saline before the surgery, and monitor closely the animals after the surgery. As for the pancreatic enzyme leakage, ensure that the organ is well cauterized during the resection step as it can rapidly cause digestive enzymes leakage in the peritoneum cavity (ascites buildup), which would result in an early humane endpoint for the animal. Providing softened food or a gel diet on the cage floor may become necessary if the recovery is lengthy.

Problem 7

Infection or inflammation of the surgical wound (Step 8 and Step 20).

Potential solution

Performing surgery with non-sterile surgical tools or using non-sterile surgical techniques may cause wound infections. Ensure the use of appropriate aseptic tools and techniques throughout the procedure to minimize the risk of infection. Improper wound closure may also allow soiled mice bedding to enter the wound, which could also cause infection. Inspect the wounds daily for 2–3 days after the surgery to either close it properly, administer additional antiseptic, or administer antibiotics as per your institution’s UACC guidelines.

Problem 8

No recurrence of the disease in the controls when it is expected (Step 25).

Potential solution

Although some variation is expected in disease recurrence after the resection surgery, no tumor recurrence in surgery-only controls could indicate that the surgery was performed too early before dissemination of the KPC cells to other organs. Delaying the surgery until the tumors get slightly bigger should be more biologically relevant for this KPC resection model. An in vivo growth curve kinetic experiment (Figure 1F) may be necessary to account for the variability of newly purchased cells. Contact L.H.T for more details.

Problem 9

High variability between different users of the protocol (Step 8 and Step 20).

Potential solution

Variability between users of this protocol is very probable as there is a lack of standardized training, and differences in surgical skills may explain these differences. All users of this protocol should perform multiple practice surgeries before experimental work. While this surgery procedure is very cumbersome, carefully planning for attrition in sample size calculations prior to starting experiments will prevent data variability, especially for survival.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Lee-Hwa Tai (lee-hwa.tai@usherbrooke.ca).

Technical contact

Technical questions on executing this protocol should be directed to and will be answered by the technical contact, Lee-Hwa Tai (lee-hwa.tai@usherbrooke.ca).

Materials availability

This study did not generate new unique reagents or materials.

Data and code availability

This study did not generate datasets or code.

Acknowledgments

We acknowledge the Craig’s Cause Pancreatic Cancer Society (Faculty Research Grant to L.-H.T.) for providing the funds to conduct this research and the Cancer Research Society for providing a doctoral scholarship to J.L.L. We would also like to thank the technical staff of the Université de Sherbrooke Animal and Transgenesis Facilities for their support of this research. The graphical abstract was created using BioRender.

Author contributions

J.L.L., M.F.M.P., and L.D. performed the experiments and collected and analyzed the data. J.L.L., N.A., S.M., and H.G. worked to troubleshoot and develop this protocol. J.L.L. wrote the manuscript. Y.C. and L.-H.T. conceptualized and supervised the study and reviewed the manuscript. L.-H.T. obtained funding for the study.

Declaration of interests

The authors declare no competing interests.