Immunophenotyping of Orthotopic Homograft (Syngeneic) of Murine Primary KPC Pancreatic Ductal Adenocarcinoma by Flow Cytometry

Summary

The experimental procedure on the immunophenotyping of murine orthotopic PDAC homografts aims at profiling the tumor immuno-microenvironment. Tumors are orthotopically implanted via surgery. Tumors of 200–600 mm³ in size were harvested and dissociated to prepare single-cell suspensions, followed by multi-immune marker FACS analysis using different fluorescently-labeled antibodies.

Abstract

Homograft (syngeneic) tumors are the workhorse of today's immuno-oncology (I/O) preclinical research. The tumor microenvironment (TME), particularly its immune-components, is vital to the prognosis and prediction of treatment outcomes, especially those of immunotherapy. TME immune-components are composed of different subsets of tumor-infiltrating immune cells assessable by multi-color FACS. Pancreatic ductal adenocarcinoma (PDAC) is among the deadliest malignances lacking good treatment options, thus an urgent and unmet medical need. One important reason for its non-responsiveness to various therapies (chemo-, targeted, I/O) has been its abundant TME, consisting of fibroblasts and leukocytes that protect tumor cells from these therapies. Orthotopically implanted PDAC is believed to more accurately recapture the TME of human pancreatic cancers than conventional subcutaneous (SC) models.

Homograft tumors (KPC) are transplants of mouse spontaneous PDAC originating from genetically engineered KPC-mice (Kras^G12D/+/P53^-/-/Pdx1-Cre) (KPC-GEMM). The primary tumor tissue is cut into small fragments (~2 mm³) and transplanted subcutaneously (SC) to the syngeneic recipients (C57BL/6, 7–9 weeks old). The homografts were then surgically orthotopically transplanted onto the pancreas of new C57BL/6 mice, along with SC-implantation, which reached tumor volumes of 300–1,000 mm³ by 17 days. Only tumors of 400–600 mm³ were harvested per approved autopsy procedure and cleaned to remove the adjacent non-tumor tissues. They were dissociated per protocol using a tissue dissociator into single-cell suspensions, followed by staining with designated panels of fluorescently-labeled antibodies for various markers of different immune cells (lymphoid, myeloid and NK, DCs). The stained samples were analyzed using multi-color FACS to determine numbers of immune cells of different lineages, as well as their relative percentage within tumors. The immune profiles of orthotopic tumors were then compared to those of SC tumors. The preliminary data demonstrated significantly elevated infiltrating TILs/TAMs in tumors over the pancreas, and higher B-cell infiltration into orthotopic rather than SC tumors.

Introduction

Pancreatic ductal adenocarcinoma (PDAC) causes nearly half a million mortalities world-wide annually, one of the top 5 cancer killers. There are few effective treatment options and no approved immunotherapies; therefore, new treatments are desperately needed. Cancers are increasingly being recognized as immunological diseases, including PDAC, in addition to the genetic diseases as known today. Immunological and genetic factors would likely determine disease prognosis as well as treatment outcomes. Tumors evade host immune surveillance and eventually advance to cause death. Many of these immune processes occur within the tumor microenvironment (TME)^1,2,3,4 where different types of immune cells interact with tumor cells, with each other and with other tumor stromal components, directly or indirectly via cytokines which ultimately determine disease outcome. Therefore, characterization of the tumor immune components of the TME, or tumor immunophenotyping, including subtyping, numeration and localization of different lineages of immune cells, is critical to understanding anti-tumor immunity. In the case of PDAC, it has been proposed that elevated tumor-infiltrating suppressive macrophages (TAM) and B-cells have led to prevention of T-cell infiltration and/or activation and high levels of fibrosis^5,6.

The common approach to investigating immune TMEs experimentally would be using surrogate tumor preclinical animal models, mainly relevant mouse tumor models⁷, particularly mouse syngeneic (homograft) or genetically engineered mouse models (GEMM) of cancers, on the assumed similarity of mouse and human for tumors and immunity^8,9. It is understood in reality that there are inherent differences between the two species^10,11.

Transplanted mouse tumors have significant operational advantages over spontaneous tumors⁷, namely synchronized tumor development, in contrast to the parental GEMM spontaneous tumor development. Homografts of spontaneous murine tumors are considered primary tumors having never been manipulated in vitro, and mirroring original mouse tumor histo-/molecular pathology⁷, as well as possible immune profiles. These murine homografts are often considered to be "a mouse version of patient-derived xenografts (PDXs)". They therefore likely have a better translatability than conventional syngeneic cell line-derived mouse tumors¹². In particular, many homografts are derived from specific GEMM where specific human disease mechanisms, e.g. oncogenic driver mutations, are engineered, and these homografts should therefore have advantages to their clinical relevance. In particular, KPC GEMM develop mouse PDAC within 15–20 weeks of age, which morphologically recapitulates human disease with predominately well- to moderately-differentiated glandular architecture and highly enriched stroma. This model also recapitulates the most common genetic features of human PDAC, namely Kras activating mutation and P53 loss-of-function, which occur in 90% and 75% of human PDAC, respectively^5,6.

Sites of transplantation have also been suggested to play a role in model translatability. The specific surrounding tissue environment, such as a corresponding orthotopic environment, could be a niche for specific tumors to progress, as opposed to the uniform subcutaneous (SC) environments for commonly transplanted tumors. It would be of particular interest if, and/or, what difference exists between the two transplantation sites, in terms of immune-microenvironment, and the relevance to human cancer, e.g. in the case of PDAC.

One of the most important aspects of immune profiling, or immunophenotyping, is to determine tumor-infiltrating immune cells of different lineages, the numbers, relative percentage within tumors, as well as their activation states, and locations. This includes tumor-infiltratrating lymphoctyes (TILs, both T- and B-), tumor-infiltrating macrophages (TAMs), tumor-infiltrating natural killer cells (NKs) and tumor-resident dendritic cells^{3,13,14,15,16,17}, and the subcellular localization of certain cells^18,19,20, etc. Fluorescence Activated Cell Sorting (FACS) or flow cytometry is a single-cell detection technology that is commonly used to measure the specific parameters of a cell. Multi-color flow cytometry measures multiple markers on a single cell^3,4,21 and is the most commonly used method to determine the numbers and relative percentage of different subsets of immune cells, including those within tumors.

This report describes procedures for profiling tumor-infiltrating immune cells: 1) Implantation of orthotopic PDAC mouse tumor homografts, along with SC implantation; 2) tumor tissue harvest and single cell preparation via tumor dissociation; 3) flow cytometry analysis of all of the cells derived from tumors as a baseline; 4) comparison of baseline profiles of both transplantation approaches.

Protocol

All the protocols and amendment(s) or procedures involving the care and use of animals will be reviewed and approved by the Crown Bioscience Institutional Animal Care and Use Committee (IACUC) prior to the conduct of studies. The care and use of animals will usually be conducted in accordance with AAALAC (Association for Assessment and Accreditation of Laboratory Animal Care) International guidelines as reported in the Guide for the Care and Use of Laboratory Animals, National Research Council (2011). All animal experimental procedures will be under sterile conditions at SPF (specific pathogen-free) facilities and conducted in strict accordance with the Guide for the Care and Use of Laboratory Animals from different government institutions (e.g. The National Institutes of Health). The protocol will need to be approved by the Committee on the Ethics of Animal Experiments at the facility institution (e.g. institutional IACUC Committee).

1. Preparation for Tumor Transplantation

1. Animal Housing
   1. Obtain generic C57BL/6 mice from a commercial breeding vendor.
   2. House mice (5) in individual ventilated cages at the following conditions: Temperature: 20–26 °C; humidity 30–70%; lighting cycle: 12 h light and 12 h dark.
   3. Use corn cob bedding that is changed weekly.
   4. For diet, provide irradiation sterilized dry granule food during the entire study period.
   5. For water, provide animals free access to sterile drinking water.
2. Donor tumor fragment preparation
   1. Begin by monitoring body weight (BW), via weighing using a balance, and tumor volume (TV), by caliper measurement, of tumor-bearing donor mice.
   2. When TV reaches 500–1,200 mm³, euthanize the animal in a biohazard hood as per protocol.
   3. Sterilize the skin around the tumor using iodophor swabs.
   4. Surgically remove the tumor (details described in section 3: Necropsy and tumor harvest) and place the tumor in a Petri dish containing 20 mL of PBS (pre-chill the media or buffer to 4 °C prior to euthanizing animals).
   5. If there is contaminating blood, transfer the tumor into another Petri dish and wash the tumor with PBS.
   6. Cut the tumor in half, removing any extra skin, vessels, calcification and/or necrosis.
   7. Choose only intact pieces of tumor and place them into a sterile 50 mL centrifuge tube and add 20 mL of PBS then transport the tube to a separate animal room for pharmacology studies.

2. Orthotopic and Subcutaneous (SC) Engraftment

1. SC Inoculation
   1. Cut tumors into 2 mm diameter pieces using a scalpel, putting 1 chunk into each trocar, for SC implantation or for later orthotopic implantation (see below).
   2. Anesthetize the recipient animal with 5% isoflurane, which is maintained by a nose cone at 1%. The animal will begin to relax, losing their righting reflex and eventually become immobile. At this depth of anesthesia they can easily be roused by painful stimuli; allow anesthesia to deepen until such responses to pain are absent.
   3. Once anesthetized, fix the mice on an experiment board in the right lateral position. Sterilize the mouse using iodophor swabs, particularly the areas surrounding the tumors.
   4. With a scalpel, make a 0.5 to 1.0 cm skin incision on the left flank, just cranial to the hip.
   5. Tunnel under the skin towards the forelimb, two to three centimeters, with blunt forceps.
   6. Aseptically transfer one cube of tumor from the medium and place deep inside the subcutaneous tunnel.
   7. Visually confirm that the tumor is deep in the tunnel (and not at the skin incision).
   8. Close the wound with wound clips.
   9. Monitor the animals post inoculation until they regain sufficient consciousness to maintain sternal recumbence. Return animals back to their cage only after their full recovery from anesthesia. Complete the Anesthesia Log, and initiate the Animal Health Chart. Weigh recipient animals daily for three days
   10. For a standard procedure, inoculate 5-10 mice per group for tumor growth monitoring.
2. Pancreatic orthotopic implantation
   1. Anesthesia and Analgesia
      1. Use a 2 mL ketamine injection and 0.42 mL xylazine injection (20 mg/mL) mixed in 5.91 sterile injection water or saline at a dose volume of 0.06–0.1 mL/20-25 g body weight.
         NOTE: According to animal welfare, analgesia is necessary both pre and post operation. 0.05–0.1 mg buprenorphine /kg, SC. The first dose is pre operation and then dosed 3 times every 4 hours post operation continually.
   2. Surgical operation for orthotopic implantation
      1. Anesthetize mice via intramuscular injection (IM) per Step 2.2.1.
      2. After the animals are fully anesthetized, fix the mice on an experiment board in the right lateral position.
      3. Keep the mice in the right lateral position. Disinfect the skin around the spleen with iodine then de-iodinate with 75% ethyl alcohol.
      4. Find the medium point of the spleen and make a 1 cm vertical incision on abdomen to expose the spleen.
      5. Draw out a part of pancreas tissue under the spleen gently with flat-tip tweezers, and suture a mouse homograft tumor piece from seed mouse on the pancreas of recipient mouse by 9-0 Absorbable surgical suture.
      6. Close the abdomen with a 6-0 silk suture by double seam. Achieve homeostasis by compression.
      7. After finishing tumor implantation, if neither bleeding nor tumor tissue leakage occurs, keep the animals in a warm cage.
      8. Monitor the animal until it regains sufficient consciousness to maintain sternal recumbency; return the animal to the animal room after full recovery from the anesthesia. Monitor the tumor bearing mice by palpating the abdomen near the spleen and select out the mice bearing orthotopic tumors.
3. Tumor-Bearing Mice Health Monitoring
   1. Check the water and food consumption daily.
   2. Examine the mouse appearance for an ungroomed hair coat, lumps, thinness, abnormal breath or ascites.
   3. Palpate the abdomen to check if there are spontaneous tumors on the liver or spleen.
   4. Weigh mice weekly using a balance.
   5. If any of the following clinical signs are observed, the mice are sacrificed for sample collection and necropsy: BW loss >20%; impaired mobility (not able to eat or drink); unable to move normally due to significant ascites and enlarged abdomen; effort in respiration.

3. Necropsy and Tumor Harvest

1. Inspect visually and by palpation for the presence of palpable tumors before termination.
2. For termination, first euthanize mice as per approved protocol, and open the abdomen and visually examine the pancreas for tumors.
3. Cut off the tumor by post mortal surgery and add it to cold PBS. Place all animals in a clean, uncharged, translucent euthanasia chamber with a special lid with a port for delivery of the carbon dioxide to be used.
4. Discharge gas into the chamber at a flow rate that produces rapid unconsciousness with minimal distress to the animal. The optimum flow rate for CO₂ should be around 2–2.5 L/min.
5. Visually observe each animal during the euthanasia procedure to assure all animals receive adequate gas concentrations and do not regain consciousness during the euthanize procedure.
6. Maintain gas flow for at least 1 min after apparent clinical death to minimize the possibility that an animal may recover (if apneic but not dead).
7. Remove adjacent non-tumor tissues. Cleaned tumor can be photographed quickly.
   Put the tumor tissues into RPMI-1640 and keep on ice before dissociation.
   NOTE: Animal carcasses are bagged and stored in the appropriate freezer to await removal.

4. Tumor Tissue Dissociation and Single Cell Preparation

1. Reagent preparation
   NOTE: The Tumor Dissociation Kit contains 2 vials of Enzyme D (lyophilized powder), 1 vial of Enzyme R (lyophilized powder), 1 vial of Enzyme A (lyophilized powder) and 1 mL of Buffer A.
   1. Prepare Enzyme D by reconstituting the lyophilized powder in each vial with 3 mL of RPMI-1640. Prepare aliquots of an appropriate volume to avoid repeated freeze-thaw-cycles.
   2. Prepare Enzyme R by reconstituting the lyophilized powder in the vial with 2.7 mL of RPMI 1640. Prepare aliquots of an appropriate volume to avoid repeated freeze-thaw-cycles.
   3. Prepare Enzyme A by reconstituting the lyophilized powder in the vial with 1 mL of Buffer A supplied with the kit. Do not vortex. Prepare aliquots of an appropriate volume to avoid repeated free-thaw-cycles.
   4. Prepare the Enzyme mix by adding 2.35 mL of RPMI-1640, 100 µL of Enzyme D, 50 µL of Enzyme R and 12.5 µL of Enzyme A into each gentleMACS C-tube.
2. Tumor Dissociation
   1. For each tumor prepare one C-tube with tumor digestion mix, please refer to section 4.1 for digestion buffer dilution and preparation.
   2. Use 3 mL digestion buffer for tumor fragments <0.8 g, and ensure the tumor is fully digested.
   3. Label with the study code, tumor, mouse ID, treatment group, and tumor weight.
   4. Collect the tumor from the mouse, wash the tumor in cold PBS and clean out the tissue attached on the tumor (such as blood vessels, fat, fascia, etc.).
   5. Put the tumor in digestion media in one well of a sterile 6 well plate.
   6. Hold the tumor in place with sterile tweezers/forceps and slice with a scalpel. Slice the tumor well enough to break into smaller pieces (~1 mm³).
   7. Place the tumor pieces back into the C-tube and use the remaining digestion buffer to wash the plate and then transfer the fluid into the C-tube that is placed on ice until digestion.
   8. Switch on the dissociator with heaters.
   9. Place tumor dissociation C-tubes upside down into sleeves of the vacant positions and adjust the status of tube positions from Free to Selected. Choose a dissociation program (37_c_m_TDK_1), followed by the selection of the required folder, where the list of Programs is displayed.
   10. After termination of the program, take C-tubes off the dissociator and spin briefly (300 x g, 4 °C) to pellet the sample.
   11. Re-suspend samples and put into a cell strainer above a 50 mL tube. Wash the cells through the cell strainer with 10 mL of wash buffer to provide a single-cell suspension.
   12. Centrifuge the tubes at 300 x g for 5 min, discard the supernatant and re-suspend the cells with 5 mL of wash buffer, count viable cells using trypan blue and/or by cell counter and adjust the cell concentraton to 1 x 10⁶ cells per tube or per sample. Include the correct isotype control antibodies to ensure staining is specific

5. Immune Panel Design and Flow Data Acquisition

1. Panel design
   1. Please see Table 1.
2. Immunostaining
   1. Fc-Block sample cells: re-suspend the cells in 200 µL of staining buffer with 1 µg/mL Fc-Block, followed by incubation on ice or 4 °C refrigeration for 15 min in the dark.
   2. Stain cells using the desired antibody/fluorescence panels (e.g. T-cell panel, macrophage panel, etc.): Add the antibody mixture diluted in Fc blocking buffer to each sample, stain for at least 30 min on ice in the dark.
   3. Add 1 mL of ice cold PBS to each tube and re-suspend the cells gently, followed by centrifugation at 300 x g for 5 min. Discard the resulting supernatant.
      1. Repeat to wash the cells twice.
   4. Stain for intracellular markers if needed, following steps 6-10, otherwise jump to step 10.
   5. Re-suspend the cell pellet by pulse vortex and add 200 µL of prepared Fixation/Permeabilization working solution for each sample. Pulse vortex again, and then incubate at 4 °C overnight (preferred) or 30 min at room temperature in the dark.
   6. Spin down the cells and remove the supernatant.
   7. Wash twice by adding 1 mL of 1x Permeabilization Buffer (made from 10x Permeabilization Buffer, diluted with distilled H₂O) followed by centrifugation and decanting of supernatant.
   8. Add intracellular marker antibody in 1x Permeabilization Buffer and incubate at room temperature for 30 min in the dark.
   9. Wash cells twice with 1 mL of 1x Permeabilization Buffer. Centrifuge and decant supernatant.
   10. Resuspend cells in 150 µL of Staining Buffer and analyze on a cytometer. Due to the fixation and permeabilization procedure, the FSC (forward-light scatter)/SSC (sidelight scatter) distribution of the cell population will be different to live cells. Therefore, the gate and voltages will need to be modified.
3. FMO Controls (Fluorescence Minus One Control)
   NOTE: Multi-color flow analysis is particularly important for the analysis of tumor-infiltrating immune cells. Therefore, there is a need to find a way to identify and gate cells in the context of data spread due to the multiple fluorochromes in a given panel. FMO (Fluorescence Minus One) Control is an important approach for this purpose.
   1. To this end, include additional mice in each group for FMO controls (at least 2 per Rx) and process individually for each tissue. After dissociation, pool tissues. For example, in a study with 4 Rx groups, 8 additional tumors should be processed individually and then pooled into one sample for FMOs.
4. Flow Instrument Setup
   1. Make compensation beads while the machine is warming up (at least 20 min).
   2. Use CS&T beads to check performance.
   3. Voltage and compensation settings: use UltraComp beads, vortex the Comp beads thoroughly before use.
   4. Label a separate 12 x 75 mm² sample tube for each fluorochrome-conjugated antibody.
   5. Add 100 µL of staining buffer to each tube. Add 1 full drop (approximately 60 µL) of the beads to each tube.
   6. Add antibodies and perform the staining procedure exactly as the sample process stated in section 4.
   7. Add 0.5 mL of staining buffer each, to completely re-suspend bead pellets via vortex.
   8. Set flow cytometer PMT voltage per target tissue for the given experiment.
   9. Run through flow cytometer for data acquisition, by gating on the singlet bead population per FSC and SSC readings.
   10. Set Flow rate around 200–300 events/s.
   11. Set appropriate compensation for a given fluorescein [FITC]-conjugated antibody, use an FL1 vs. FL2 dot plot.
   12. Place a quadrant gate so that negative beads are within lower left quadrant and the positive beads are in the upper or lower right quadrant. Adjust the compensation values until the median fluorescence intensity (MFI) of each population (as shown in the quadrant stats window) is approximately equal (i.e. for FL2-%FL1, the FL2 MFI of both beads should be similar).
   13. Repeat steps 5.4.11 and 5.4.12 for all tubes.
   14. Proceed to acquiring the actual stained samples. Run the compensation wizard and save the settings with the format "date experiment your initials".

6. Flow Data Analysis and Presentation

1. Analyze data by Flowjo and/or Kaluza.

Results

Orthotopic implantation of PDAC resulted in rapid tumor growth similar to that seen for SC implantation. After the donor tumor fragments were implanted into recipient mice, both subcutaneously and orthotopically according to the protocols described in Steps 2.1 and 2.2, the implanted KPC homograft tumors demonstrated similar rapid growth as shown in Figure 1A. KPC homograft tumors harvested at different time points are shown in Figure 1B<...

Discussion

Although studies using SC tumors are more readily conducted, orthotopically implanted tumors models can potentially be more relevant for preclinical pharmacology studies (particularly I/O investigations) to provide enhanced translatability. This report aims at helping the interested readers/audience to be able to directly visualize the technical procedures that can be used in their respective research. Our protocols demonstrate that orthotopic implantation of PDAC can result in efficient tumor growth, similar to SC impla...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anesthesia machine	\	SAS3119	\
Trocar 20	\	\	2 mm
Petri dish	\	\	20 mm
100x antibiotic and antimycotic	\	\	\
Iodophor swabs	\	\	Daily pharmacy purchase
Alcohol swabs	\	\	Daily pharmacy purchase
Liquid nitrogen	Air chemical	\	\
Biosafety hood	AIRTECH	BSC-1300IIA2	\
FACS machine LSRFortessa X-20	BD	LSR Fortessa	\
antibodies	BD	\	\
Trevigen MD or BD Matrigel Basement Membrane Matrix High concentration	BD	354248	\
FACS buffers	BD	554656	Mincing buffer
Brilliant Staining Buffer	BD	563794	\
Mouse BD Fc Block	BD	553142	\
cell filters	BD-Falcon	352350	70 µm
routine blood tube	BD-Vacutainer	365974	2 mL
Kaluza	Beckman		vs 1.5
6-well plates	Corning	3516	\
Foxp3 Fix/Perm kit	ebioscience	00-5523-00	\
UltraComp eBeads	ebioscience	01-2222-42	\
Centrifuge	eppendorf	5810R,5920R	\
FlowJo software	FlowJo LLC	\	vs 10.0
PBS	Hyclone	SH30256.01	50 mL
RPMI 1640	Hyclone	SH30809.01	\
Disposable, sterile scalpels	Jin zhong	J12100	11#
knife handle	Jin zhong	J11010	\
eye scissors and tweezers	Jin zhong	Y00030	Eye scissors 10 cm
eye scissors and tweezers	Jin zhong	JD1060	Eye tweezers 10 cm with teeth
Portable liquid nitrogen tank	Jinfeng	YDS-175-216	\
Electronic balance	Metter Toledo	AL204	0-100 g
Miltenyi C-tubes	Miltenyi	130-096-334	\
Miltenyi Gentle MACS with heater blocks	Miltenyi	120-018-306	\
Tumor Dissociation Kit	Miltenyi	130-096-730	\
Cell counter	Nexcelom	Cellometer	Cellometer Auto T4
cryopreservation tube	Nunc	375418	1.8 mL
Cultrex High Protein Concentration (HC20+) BME	PathClear	3442-005-01	\
syringes	Shanghai MIWA medical industry	\	1-5 mL
Studylog software	Studylog	\	software
Studylog-Balance and supporting USB	OHAUS	SE601F	Balance and supporting USB
Studylog-Data line of vernier calipers	Sylvac	926.6721	Data line of vernier calipers
Caliper	Sylvac	910.1502.10	Sylvac S-Cal pro
Sterilized centrifuge tubes	Thermo	339653	50 mL
Sterilized centrifuge tubes	Thermo	339651	15 mL
Ice bucket	Thermo	KLCS-288	4 °C
Ice bucket	Thermo	PLF-276	-20 °C
Ice bucket	Thermo	DW-862626	-80 °C
RNAlater	Thermo	am7021	\