Intradermal Tumor Implantation as a Robust Model for Murine Cancer Intratumoral Immunotherapy Studies

Summary

With increasing interest in intratumoral immunotherapy, selecting tumor models that are accessible and practical is essential. Subcutaneous tumors are easy to establish but difficult to visualize and manipulate. In this protocol, we describe methods for generating intradermal tumors that provide improved visibility and accessibility.

Abstract

Establishing reproducible murine tumor models is essential for evaluating cancer biology and immunotherapy strategies. Intratumoral immunotherapy treats established tumors with immune-stimulating reagents to reverse local immune suppression and stimulate systemic antitumor immunity. Most cancer studies in mice establish the tumors in the subcutaneous space, which has inherent challenges for experimental intratumoral immunotherapy due to the inability to visualize intratumoral injections. Intradermal injection of tumor cells provides a reliable approach to generate visualizable tumors that support accurate and reproducible intratumoral injection and monitoring of growth kinetics and therapeutic responses. Here, we present a reproducible methodology for establishing and treating intradermal tumors using the murine B16F10 melanoma model in C57BL/6 mice as our example. Tumor cell suspensions are injected intradermally, and 7–10 days after tumor implantation, once tumors reach ~60 mm³, an intratumoral injection of immune-stimulating agents or a physical treatment like electroporation or heating can be directly applied. This allows precise delivery of treatments and measurement of tumor growth with calipers and avoids the challenge of the inherent variability of multiple treatments of subcutaneous tumors that cannot be visualized. By providing detailed insight into this technique, this article aims to support reproducibility and advance pre-clinical research in intratumoral immunotherapy.

Introduction

Murine tumor models constitute an indispensable tool in cancer research. In an optimal mouse model, it would be possible to induce tumors that closely mimic human tumors at both genetic and morphologic levels¹. This enables the assessment and efficiency of novel therapeutic approaches in different fields, including intratumoral immunotherapy (ITIT). ITIT is an emerging field in cancer treatment that consists of directly administering immune-stimulating agents into the tumor micro-environment (TME). This localized delivery can enhance local immune activation, reverse the immunosuppressive tumor micro-environment, and trigger systemic antitumor immune responses, known as the abscopal effect². However, the efficiency, accuracy, and consistency of intratumoral administration depend on the ability to visualize and control injection parameters in different tumor models.

Most pre-clinical cancer studies use subcutaneous tumor implantation. However, this approach has limitations for intratumoral therapies because the tumor cannot be directly visualized during injection unless surgery is performed. Many intratumoral immunotherapy strategies require precise delivery of reagents into small mouse tumors and often entail accomplishing that task multiple times. Without visual confirmation, injection accuracy and reagent retention within subcutaneous tumors become poorly controlled variables. This could result in misinterpretation/variation of experimental data³.

Intradermal tumor implantation provides a direct, accessible, visualizable, and reproducible alternative for tumor observation, measurement, and intratumoral injection, thus enhancing consistency. Moreover, studies have shown that tumors implanted intradermally, as opposed to subcutaneous injections, enhance immunogenicity and suppress the tumorigenicity of tumor cells⁴. Intradermally injected tumors induced rapid and strong dendritic cell trafficking to lymph nodes, resulting in earlier and more robust activation of tumor-specific cytotoxic T cells⁴. The present study describes a detailed and reproducible protocol for establishing intradermal B16F10 melanoma tumors in C57BL/6 mice and performing subsequent intratumoral treatments.

Protocol

All procedures were conducted with the approval of, and in compliance with, the Institutional Animal Care and Use Committee (IACUC) guidelines⁵. All animal procedures were performed in Dartmouth’s Animal Facility. Female C57BL/6 mice, 6–8 weeks of age, were housed under standard conditions on ventilated racks with food and water. The reagents and the equipment used are listed in the Table of Materials. 

1. Shaving mice

NOTE: All personnel must complete institutional training in live animal handling and anesthesia prior to performing these procedures.

1. Prior to intradermal injection, shave the flank of each mouse as described below; the right flank was used for injections and the described procedures. Shave the mice the day prior to injecting tumor cells.
2. Position the mouse on a stable surface. Scruff the mouse by the skin of the nape with the left thumb and index finger, and gently tuck the left side of the body into the palm to expose the right side. Use the ring and/or pinky fingers to hold the left leg and tail, ensuring the right flank is fully exposed.
3. Shave the fur from the hind leg up to the lower ribs to create a clear area for injection.
4. Both flanks may be shaved if additional practice space is needed.
   NOTE: Beginners can anesthetize mice when learning to shave and properly scruff.

2. Practice injections with dye

NOTE: Practice can be performed on both live (anesthetized) and euthanized mice, and on either sex. Male mice typically have thicker and less pliable skin compared to females, making the procedure more technically challenging. For this reason, practicing on males is useful for developing proficiency.

1. Mix food coloring with phosphate-buffered saline (PBS) in a 1.5 mL Eppendorf tube until the solution appears dark (blue food coloring was used in this study).
2. Obtain practice mice and anesthetize them in accordance with institutional animal care guidelines. Apply ophthalmic ointment to both eyes to prevent corneal drying.
3. Place the anesthetized mouse under a nose cone and tilt the body to expose the right flank. Confirm adequate anesthesia by gently pinching the footpad and observing no response.
4. Load 30 µL of dye solution into a 0.3 mL insulin syringe with a 29 G, ½” needle. Remove all air bubbles.
5. Using the non-dominant hand, gently pinch and stretch the leg/skin to create a smooth, taut surface (avoid lifting the skin upward). Insert the needle bevel-up into the dermis at a shallow parallel angle to the skin. Keep the needle visible beneath the surface to confirm dermal placement.
   NOTE: During practice, test different hand grip positions to maintain stable skin tension while keeping fingers clear of the needle path. Insert the needle ~2 mm into the skin. To reduce backflow, insert the needle ~3 mm, then withdraw slightly to create a “pocket” before injection. This helps retain the dye at the injection site.
6. Slowly inject all 30 µL of the dye solution until a small, distinct “bubble” forms under the skin surface
   NOTE: The absence of visible bubbles suggests that the injection was delivered subcutaneously rather than intradermally. Maintaining the bevel in an upward position helps ensure accurate dermal placement and prevents overly deep injection.
7. Continue practicing until consistently forming intradermal bubbles. Up to six to eight injections can be performed per side.
8. After completing practice injections, euthanize the mice in accordance with approved Institutional Animal Care and Use Committee (IACUC) protocols⁵.

3. Preparation of tumor cells for injection

NOTE: Perform all cell-related steps in a sterile hood. Be mindful that different murine tumor lines grow differently.

1. Thaw tumor cells (B16F10 used in this study) several days before intradermal injections.
2. Culture cells to 70%–80% confluence while maintaining adequate fresh media
   NOTE: Avoid 100% confluence, which halts proliferation and impairs tumor growth.
3. Harvest and prepare cells at 7 x 10⁶ cells/mL in either 1x PBS, serum-free RPMI, or other media used for growing the cells.
   NOTE: The B16F10 tumor model used in the lab uses 2 x 10⁵ cells in 30 µL. The 30 µL injection volume is appropriate, but the number of cells can be adjusted as needed to establish the desired tumor model. Cells will settle and collect at the bottom of the tube, so to ensure the concentration of cells is constant, periodically gently mix the cells to keep them suspended.
4. Keep cells on ice after processing and proceed quickly to inject. Prolonged storage on ice reduces cell viability, which can decrease the number of live cells delivered and negatively impact tumor establishment⁶.
   NOTE: Cells are harvested and used right away in the lab. Injections are done within 20–60 min of processing the cells and putting them on ice. Use a partner when working with large cohorts to minimize time on ice.

4. Intradermal injections

1. Anesthetize experimental mice. Apply ophthalmic ointment to both eyes to prevent corneal drying.
   NOTE: Anesthesia with isoflurane gas is used by the lab, but injectable anesthetics can be used.
2. Following anesthesia induction, place the mouse under a nose cone and position the body to fully expose the right flank.
3. Gently invert the cell suspension to ensure even mixing.
4. Draw 30 µL of cell suspension into a 0.3 mL syringe with a 29 G, ½” needle. Remove air bubbles.
5. Using the non-dominant hand, gently stretch the leg/skin to the side to make it smooth and tight (do not lift it upward). Insert the needle bevel-up into the dermis at a shallow parallel angle. The needle should stay visible under the skin, showing it’s not too deep.
   NOTE: The needle depth should be ~2 mm into the skin. To avoid backflow of the cells, insert the needle ~3 mm in the skin, pull the needle back slightly to create a “pocket” for the cells to be injected into. This reduces backflow significantly.
6. Very slowly inject all 30 µL of cell suspension until a small, visible “bubble” forms under the skin. Wait 5–10 s before removing the needle.
   NOTE: Removing the needle too quickly can cause the solution to come out of the skin due to the pressure.
7. Repeat for each mouse. Replace syringes after each cage (4–5 mice) to prevent needle dulling.
8. Monitor all mice until they have fully recovered from anesthesia.
9. Monitor the mice for tumor establishment and overall health.

5. Intratumoral injections

NOTE: Tumors are typically ready for treatment 7–10 days post-implantation, once they reach the appropriate size. Appropriate size used in the lab is ~50 mm³ or higher using the formula: Volume = (Length/2) * (Width*Width)^7,8.

1. Prepare all treatment solutions in advance (e.g., 1x PBS was used in this study). Keep solutions on ice or at room temperature as required by the specific treatment.
2. Induce anesthesia according to institutional guidelines.
3. Place the mouse under a nose cone to maintain anesthesia, and position it so the tumor is exposed.
4. Draw the required volume (e.g., 30 µL used here) into a 0.3 mL insulin syringe with a 29 G, ½” needle. Remove air bubbles.
   NOTE: Volume will differ based on the treatment of choice and optimized conditions.
5. Insert the needle horizontally into the tumor and advance toward the center of the tumor mass.
   NOTE: Avoid inserting from the top or at a steep angle, as this can result in the injection being delivered too deep or into surrounding tissue. The treatment described was administered as a single injection. The doses and frequency are all dependent on the treatment of choice and optimized conditions. With small tumors, a single dose is adequate in distributing the treatment, but with larger tumors, multiple injections may be needed to distribute the treatment equally through the tumor mass.
6. Inject the treatment slowly and steadily. Pause for 10 s before slowly withdrawing the needle to minimize backflow or leakage. Some tumors have high internal pressure, and care should be taken to ensure the injection does not leak out through the needle hole.
7. Repeat the procedure for each mouse. Replace syringes after each cage of 4–5 mice and when switching treatment groups to prevent cross-contamination.
8. Monitor mice post-anesthesia until they regain mobility and normal respiration. Continue daily monitoring for treatment response and overall health.
   NOTE: Be mindful that different cell lines may cause tumors to ulcerate at different times. Each experiment and each IACUC protocol has clearly stated endpoints and exclusion criteria. In the lab, endpoints for mouse experiments are 1500 mm³ and ulcerations that occur are accepted and not excluded from the study.

Results

When practicing injections with dye, successful intradermal delivery can be verified without euthanizing the mouse by observing the formation of a visible “bubble” at the injection site. A properly performed intradermal injection produces a large, well-defined, and raised “bubble” (Figure 1A–F). Small or flat bubbles suggest that the injection may be partially subcutaneous. If no “bubble” forms and the dye or cells spread beneath the skin, the injection was likely subcutaneo...

Discussion

The success of this protocol (Figure 4) depends on maintaining a consistent intradermal injection depth and precise needle placement to ensure delivery into the dermis rather than the subcutaneous layer. If the needle angle is too steep or the injection pressure is too high, the dye or tumor cells may be deposited through the skin into the subcutaneous space. After injection, pausing briefly before slowly withdrawing the needle helps prevent leakage or formation of doublet tumors (“sno...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1.5 mL Natural Centrifuge Tube	USA Scientific	1615-5500
150 cm2 Cell Culture Flask	Corning	430825
Fluriso	Vet One	13985-528-60	isoflurane
Food Coloring	Durkee		for practice
Insulin syringe 0.33 x 12.7 mm	Exel	26018
Mini Arco	Wahl	08787-450A	clippers to shave the mice
PBS, 1X w/o calcium & magnesium	Corning	06325002	pH 7.4 +/- 0.1
RPMI 1640, 1x	Corning	12225004
SomnoFlo Low-Flow Electronic Vaporizer	Kent Scientific	SF-01	anesthesia machine