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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

We present a method for studying tumor cell redissemination from lung metastases involving a surgical protocol for the selective photoconversion of lung metastases, followed by the identification of redisseminated tumor cells in tertiary organs.

Abstract

Metastasis - the systemic spread of cancer - is the leading cause of cancer-related deaths. Although metastasis is commonly thought of as a unidirectional process wherein cells from the primary tumor disseminate and seed metastases, tumor cells in existing metastases can also redisseminate and give rise to new lesions in tertiary sites in a process known as "metastasis-from-metastases" or "metastasis-to-metastasis seeding." Metastasis-to-metastasis seeding may increase the metastatic burden and decrease the patient's quality of life and survival. Therefore, understanding the processes behind this phenomenon is crucial to refining treatment strategies for patients with metastatic cancer.

Little is known about metastasis-to-metastasis seeding, due in part to logistical and technological limitations. Studies on metastasis-to-metastasis seeding rely primarily on sequencing methods, which may not be practical for researchers studying the exact timing of metastasis-to-metastasis seeding events or what promotes or prevents them. This highlights the lack of methodologies that facilitate the study of metastasis-to-metastasis seeding. To address this, we have developed - and describe herein - a murine surgical protocol for the selective photoconversion of lung metastases, allowing specific marking and fate tracking of tumor cells redisseminating from the lung to tertiary sites. To our knowledge, this is the only method for studying tumor cell redissemination and metastasis-to-metastasis seeding from the lungs that does not require genomic analysis.

Introduction

Metastasis is the leading cause of cancer-related deaths1.Β Metastatic cancer arises when cells from the primary tumor disseminate throughout the body and proliferate into clinically detectable tumors in distant organs2,3.

Although metastasis is commonly thought of as a unidirectional process wherein tumor cells disseminate from the primary tumor and colonize distant organs4, increasing clinical and experimental evidence suggests that a more complex, multi-directional process is at play. It has been shown that circulating tumor cells can reseed the primary tumor (if still in place)5,6,7,8,9, and tumor cells from existing metastatic foci can travel to tertiary sites and give rise to new lesions10,11,12,13. Indeed, evidence from recent genomic analyses suggests that some metastatic lesions arise not from the primary tumor, but from other metastases-a phenomenon known as "metastasis-from-metastases" or "metastasis-to-metastasis seeding"14,15,16. Metastasis-to-metastasis seeding can perpetuate the disease process even after removal of the primary tumor, increasing metastatic burden, and decreasing patient quality of life and survival. Therefore, understanding the processes behind metastasis-to-metastasis seeding is crucial to refining treatment strategies for patients with metastatic disease.

Despite the potentially severe clinical implications, little is known about metastasis-to-metastasis seeding, due in part to logistical and technological limitations. Human studies are limited by a paucity of clinical samples. Clinical resection and biopsy of metastatic lesions are uncommon, as is the biopsy of seemingly healthy organs, where single disseminated tumor cells may lurk. This means that human studies are typically only possible using autopsy samples from individuals whose primary tumors are either still in place or were previously resected but are still available to researchers. When such samples are available, lineage analyses of cancer progression must be performed using sequencing methods14. However, bulk sequencing of matched primary tumors and metastases does not have the sensitivity needed for comprehensive lineage tracing. For instance, bulk sequencing of one lesion may reveal a subclone that is undetectable in any of its matched lesions. In this case, one would be unable to determine the origin of this subclone. It may have been present in the primary tumor or another metastasis at a frequency below the limit of detection, or it may have arisen after the initial colonization of the metastatic lesion it was found in. Single-cell sequencing provides increased sensitivity, but its high cost limits the large-scale application of this technique. The retrospective nature of these studies also means that they provide limited insight into transient metastatic events and the disease landscape at different time points.

In animal models, recent technological advances now allow for prospective phylogenetic mapping with high spatial and temporal resolution17,18,19,20. These techniques utilize CRISPR/Cas9 genome editing to engineer cells with an evolving barcode - heritable mutations that accumulate over time. Upon sequencing, the lineage of each cell can be traced based on the mutational profile of its barcode17,18,19,20. Indeed, such technology is already being used to map metastasis-to-metastasis seeding. In a recent paper, Zhang et al. demonstrated that breast and prostate cancer cells in bone metastases redisseminate from the bone to seed secondary metastases in multiple organs21.

While these novel methods have great potential to generate detailed, high-resolution phylogenetic maps of cancer progression, they are highly impractical for those studying the exact timing of metastasis-to-metastasis seeding events and what promotes or prevents them. Filling these knowledge gaps is crucial to refining our understanding and treatment of metastatic cancer, but there is a noticeable lack of technologies to facilitate such studies. To address this need, we recently developed - and present herein - a novel technique that allows us to specifically mark tumor cells via photoconversion in a metastatic site (the lung) and subsequently reidentify them in tertiary organs. Using this technique, we recently showed that breast cancer cells do redisseminate from lung metastases and seed tertiary organs13. This technique may also be used to determine the timing of redissemination events within a narrow window and quantify redisseminated tumor cells, facilitating the study of organotropism of redisseminated cells and what promotes/prevents redissemination.

While photoconversion and locally inducible cre/lox systems that permanently replace one fluorescent protein with another have been previously used to mark and track tumor cells11,22,23, to our knowledge, no approach for spatiotemporal marking of tumor cells has been optimized to target the lung - one of the most common sites of metastasis among men and women diagnosed with any of the 14 most common cancers24. Any cancer cell type and any protocol for lung metastasis generation may be used with our procedure, making it broadly useful for metastasis researchers. All cancer cells used to generate lung metastases should express a photoconvertible or photo-switchable protein, and researchers may choose which protein to use based on their specific needs and resources. In this study, we used 6DT1 breast cancer cells that stably expressed the photoconvertible green-to-red fluorescent protein Dendra2 (6DT1-Dendra2 cells)25 tagged to the histone H2B. We injected 5.0 Γ— 104 6DT1-Dendra2 cells into the fourth mammary fat pad of female Rag2-/- mice. Primary tumors were palpable between 12 and 16 days after injection and were not resected for the duration of the experiment. Spontaneous lung metastases developed between 19 and 26 days after tumor cell injection. Photoconversion surgeries were performed between 26 and 29 days after tumor cell injection. Mice were sacrificed by 72 h post surgery due to lung metastasis burden.

Protocol

All procedures described in this protocol have been performed in accordance with guidelines and regulations for the use of vertebrate animals, including prior approval by the Albert Einstein College of Medicine Institutional Animal Care and Use Committee.

Prior to surgery, lung metastases are to be generated in mice using cancer cells that express a photoconvertible/photo-switchable protein; several protocols for lung metastasis generation have been published26,27,28.

1. Preparation for surgery

  1. Prepare distinct work areas for mouse preparation (hair removal and intubation), surgery, and recovery.
  2. Sterilize all surgical instruments in an autoclave.
    NOTE: As a tips-only technique is used for this surgery, a hot bead sterilizer may be used to resterilize instruments for subsequent procedures.
  3. Turn on the heated surgical platform and allow it to reach and stabilize at 37-40 Β°C.
  4. Induce anesthesia with 5% isoflurane, and then lower the isoflurane to 3%. Perform toe pinch tests periodically throughout the procedure to assess anesthesia adequacy.
  5. Place the anesthetized mouse in a right lateral decubitus position. Remove the hair of the upper-left chest/side body (see Figure 1A) by applying depilatory cream to the area. Dampen a piece of gauze or paper towel with water and wipe the hair and depilatory cream away. Repeat as necessary until all hair is removed from the surgical field.
    NOTE: Do not leave depilatory cream on the mouse for longer than 20 s, as this can damage the skin.
  6. Tie a double knot ~3 mm above the base of a 22 G catheter with 2-0 silk suture. Leave 2-inch-long tails.
  7. Intubate the mouse with this catheter as previously described29,30. To confirm successful intubation, attach an inflation bulb to the end of the catheter and gently squeeze.
    NOTE: Mice that have been intubated successfully will experience bilateral chest rise with bulb squeeze.
  8. Tightly secure the 2-0 silk suture around the mouse's snout with a double knot to keep the intubation catheter in place.

2. Surgery to expose the lung

NOTE: Perform all steps of the surgery (Figure 1), including photoconversion, in a hood or laminar flow cabinet to avoid contamination of the surgical field.

  1. Wash hands with antiseptic soap and don new sterile gloves.
    NOTE: As a tips-only technique is used for this surgery, non-sterile gloves may also be used. Sanitizing non-sterile gloves with an alcohol-based disinfectant is recommended.
  2. Prepare a 0.1 mg/kg dose of buprenorphine (0.03 mg/mL) and inject subcutaneously for analgesia.
    NOTE: Multimodal analgesia, including local pain blockers and nonsteroidal anti-inflammatory drugs, should be used in conjunction with buprenorphine if they are not expected to interfere with the biology of interest.
  3. Apply ophthalmic ointment to both mouse eyes to prevent corneal damage.
  4. Place the mouse in the right lateral decubitus position on the heated surgical platform.
  5. Connect the ventilator to the intubation catheter. Take care to hold the catheter steady, as even slight movements can disturb the catheter placement and lead to poor ventilation.
  6. Observe stable, ventilator-controlled, bilateral chest rise and fall.
  7. Secure the limbs to the surgical platform with labeling tape. Stabilize the surgical field by placing another piece of tape along the mouse's back and securing the dorsal skin and fur to the surgical platform (Figure 1A).
  8. Open the sterilized surgical instruments.
  9. Apply chlorhexidine solution to the mouse's skin to sterilize the surgical site.
  10. Identify the area ~7 mm to the left of the sternum and ~7 mm above the subcostal margin. Lift the skin over this area with forceps and make a ~10 mm circular incision using sharp micro-dissecting scissors, taking care to cut only the skin.
  11. Remove the soft tissue over the ribcage (Figure 1B). Cauterize any major vessels at both ends with a cautery pen.
    NOTE: In addition to excising all soft tissue underlying the 10 mm circular incision in the skin, removing some additional non-muscular soft tissue from the perimeter facilitates future steps of the procedure.
  12. With one hand, use forceps to elevate the 6th or 7th rib. With the other hand, use a single blade of the blunt micro-dissecting scissors - angled parallel to the chest wall, rounded edge facing down - and carefully pierce the 6th or 7th intercostal muscle to enter the thoracic cavity (Figure 1C). Gently rotate the scissors as needed and cut to make a ~10 mm incision in the intercostal space, taking care to avoid touching the lung tissue and cutting the ribs.
  13. Delicately discharge compressed air towards the intercostal incision to increase the space between the lung and the chest wall. Discharge the air first at one's wrist or hand to find the appropriate flow strength and clear the nozzle of any debris, then in short bursts at the incision to prevent lung injury.
  14. With one hand, use forceps to elevate the 6th or 7th rib. With the other hand, hold the retractor shut and insert it into the intercostal incision. Orient the blades of the retractor such that they are parallel to the chest wall, taking care to avoid touching the lung itself (Figure 1D).
  15. Once the retractor is in place, slowly release the handle, allowing the retractor to open and expose the lung (Figure 1E). Figure 2A shows representative images of exposed lung metastases prior to photoconversion, including what they look like to the naked eye, and in the FITC (green, non-photoconverted) and TRITC (red, photoconverted) fluorescent channels while using wide-field illumination.

3. Lung metastasis photoconversion

NOTE: Details and variations on the following steps can be found in the Discussion.

  1. Gently apply 2-3 drops of sterile PBS warmed to 37 Β°C to the exposed lung tissue to prevent drying.
  2. Place a piece of sterile aluminum foil with a small cutout over the mouse. Position the cutout directly over the exposed lung tissue and size it to expose only the open portion of the thorax and a few millimeters of surrounding soft tissue and skin to ensure photoconversion of only the exposed lung.
  3. Place a box with a cutout over the mouse and aluminum foil. Ensure that the cutout is directly over the exposed lung.
  4. Place the photoconversion lamp directly over the cutout on the box (Figure 1F).
  5. Turn on the photoconversion lamp and allow 6 min of continuous illumination of exposed lung tissue.
    NOTE: Monitor the mouse's vital signs using the physiologic monitoring programs on the ventilator.
  6. After photoconversion, turn off the lamp, and remove the lamp, box, and aluminum foil mask from the mouse.

4. Procedure to close chest wall

  1. Repeat step 2.13 to separate the lung from the chest wall.
  2. Carefully take hold of the retractor handle and squeeze to close the blades.
  3. Maneuver the closed retractor out of the intercostal space, taking care to avoid touching the lung tissue.
  4. Using the 5-0 silk suture, close the intercostal incision with a simple continuous suture pattern that incorporates the rib on both sides of the incision (Figure 1G). Pull the suture tight to ensure that the wound edges are appositional and re-establish the integrity of the chest wall (Figure 1H). Take care not to overtighten the suture as this may tear the intercostal muscles.
  5. Secure the suture by knotting the two ends of the suture together 4x. Cut the suture tails as close to the knot as possible.
  6. Close the skin with surgical staples.
  7. Remove excess air from the thoracic cavity with a 1 mL insulin syringe with a 28 G needle attached. Locate the xiphoid process tactilely, and insert the needle just below, advancing toward the left shoulder to enter the thoracic cavity through the diaphragm. Draw back on the syringe to ~1 mL; remove the needle from the mouse and discharge the air from the syringe. Repeat this process 3-4x.
    NOTE: Take care not to puncture any internal organs. Furthermore, it is important to use a 1 mL syringe and repeat the procedure 3-4x rather than using a larger volume syringe and trying to remove all the air at once. Use of a larger volume syringe may result in too great of a vacuum in the thoracic cavity and lead to lung tissue distension and damage.
  8. Turn off the isoflurane.
  9. Continue ventilation with 100% oxygen until the mouse shows signs of awakening.
  10. Once the mouse shows signs of awakening, disconnect the intubation catheter from the ventilator, cut the suture around the snout, and extubate the mouse.
  11. Untape the mouse and transfer it to a clean cage placed ~2 feet below a heat lamp. Monitor until fully recovered. If the mouse shows signs of breathing difficulty or lack of mobility, anesthetize it with 5% isoflurane for 5 min, ensure adequate anesthesia by observing a loss of reflexes upon toe pinch, and euthanize via cervical dislocation.
  12. Once the mouse fully awakens from anesthesia and begins to move, prepare another 0.1 mg/kg dose of buprenorphine and inject it subcutaneously for postoperative analgesia.
  13. Following the surgery, ensure that the mice are housed individually. Provide antibiotics in the drinking water (enrofloxacin, final concentration 0.4 mg/mL).
  14. In the 3 days following the surgery, check the mouse twice per day for signs of distress. For pain management, administer 0.1 mg/kg buprenorphine (0.03 mg/mL) subcutaneously every 4-6 hours. Euthanize the mouse if it shows signs of infection, immobility, or breathing difficulty. After the third day, mice can be checked once per day.

5. Sample processing and detection of photoconverted cells using tissue clearing

  1. Between 3 and 5 days (or the desired length of time) after photoconversion surgery, anesthetize the mouse with 5% isoflurane, reduce isoflurane from 5% to 2.5% after induction, ensure adequate anesthesia by observing a loss of reflexes upon toe pinch, and perform a terminal transcardial perfusion with 10 mL of room-temperature PBS, followed by 10 mL of 4% paraformaldehyde chilled to 4 Β°C as previously described31.
  2. Collect the organs of interest and optically clear them as previously described31.
  3. Image the cleared tissues with a light sheet microscope as previously described31. Alternatively, place the cleared tissues in a glass bottom dish and image in the FITC and TRITC channels to visualize green (non-photoconverted) and red (photoconverted) cells using a confocal, spinning disk, or multiphoton microscope.

6. Sample processing and detection of photoconverted cells using tissue disaggregation

  1. Anesthetize the mouse with 5% isoflurane for 5 min, ensure adequate anesthesia by observing a loss of reflexes upon toe pinch, and sacrifice via cervical dislocation 3-5 days after photoconversion.
  2. Collect and digest the organs of interest as previously described13.
  3. Plate the digested tissues and place them in an incubator to adhere overnight as previously described13.
  4. The following day, image the plated cells in the FITC and TRITC channels to visualize green (non-photoconverted) and red (photoconverted) cells as previously described13.

Results

The steps of the surgery described in this protocol are illustrated in Figure 1. In brief, the mouse is anesthetized, and hair is removed from the left thorax. The mouse is then intubated and ventilated, which allows the mouse to receive oxygen while the thoracic cavity is open. Soft tissue is removed to expose the ribcage, and an incision is made in the 6th or 7th intercostal muscle. A retractor is inserted in the intercostal breach and released to spread the adjacent ...

Discussion

In this paper, we describe a surgical protocol for the selective photoconversion of tumor cells in the lung. This technique enables researchers to selectively mark tumor cells in the lung and track their fate by reidentifying them throughout the body at a later time point, facilitating the study of metastasis from lung metastases. Using this protocol, it was possible to visualize photoconverted cells in the brain, liver, and non-photoconverted right lung of mice that had undergone the surgery with photoconversion, indica...

Disclosures

The authors have no conflicts of interest to declare.

Acknowledgements

The authors would like to thank Wade Koba for his assistance with micro computed tomography (S10RR029545), Vera DesMarais and Hillary Guzik of the Analytical Imaging Facility for their training and assistance with microscopy, the Einstein Montefiore Cancer Center, the National Cancer Institute (P30CA013330, R01CA21248, R01CA255153), the Gruss Lipper Biophotonics Center, the Integrated Imaging Program for Cancer Research, a Sir Henry Wellcome Postdoctoral Fellowship (221647/Z/20/Z), and a METAvivor Career Development Award.

Materials

NameCompanyCatalog NumberComments
0-30 V, 0-3 A Power SupplyMPJA9616 PS
12 VDC, 1.2 A Unregulated Plug SupplyMPJA17563 PD
28 G 1 mL BD Insulin SyringeBD329410
400 nm light emitting diode array lampLedEngin Inc.897-LZPD0UA00Photoconversion lamp, custom-built (individual parts included below)
5-0 braided silk suture with RB-1 cutting needleEthicon, Inc.774B
9 cm 2-0 silk tieEthicon, Inc.LA55G
Baytril 100 (enrofloxacin)Bayer (Santa Cruz Biotechnology)sc-362890RxAntibiotic used in drinking water
BuprenorphineHospira0409-2012-32Analgesic
Cables (Cable Assemblies) 2.1 DC JACK-STRAIGHT 72"Β  BLACK/ZIP CORDMouser172-7426-E
Cables (Cable Assemblies) 2.5 JK-ST 72" ZIP CDMouser172-0250
Chlorhexidine solutionDurvet7-45801-10258-3Chlorhexidine Disinfectant Solution
Compressed air canisterFalconDPSJB-12
Extra Fine Micro Dissecting Scissors 4" Straight Sharp/Sharp 24 mmRoboz SurgicalRS-5912Sharp Micro Dissecting Scissors
Fiber-optic illuminatorO.C. White CompanyFL3000Used during mouse intubation
Gemini Cautery KitHarvard Apparatus726067Cautery pen
Germinator 500CellPoint ScientificGERΒ 5287-120VBead Sterilizer
Graefe forcepsRobozRS-5135
High power LEDs - single color ultraviolet 90 wattsMouserLZP-D0UA00
Infrared heat lampBraintree ScientificHL-1
Isoflurane SOL 250 mL PVLCovetrus29405Anesthetic
Isoflurane vaporizerSurgiVetVCT302
Jacobson needle holder with lockKalson SurgicalT1-140
Labeling tapeFisher ScientificS68702
LED Lighting Reflectors CREE MP-L SNGL LENS REFLECTOR & LOC PINMouser928-C11395TM
Long cotton tip applicatorsMedline IndustriesMDS202055
Masscool / Soccket 478 / Intel Pentium 4/Celeron up to 3.4GHz / Ball Bearing / Copper Core / CPU Cooling FanCompUSA#S457-1023
Micro Dissecting Scissors 4" Straight Blunt/BluntRoboz SurgicalRS-5980Blunt Micro Dissecting Scissors
Murine ventilatorKent ScientificΒ PS-02PhysioSuite
Nair Hair Removal LotionAmazonB001RVMR7KDepilatory cream
Personnet mini retractorRobozRS-6504Retractor
Phosphate Buffered Saline 1xFisher Scientific14190144PBS
pLenti.CAG.H2B-Dendra2.WAddgene51005Dendra2 lentivirus
PuralubeHenry Schein Animal Health008897Eye Lubricant
Rodent intubation standBraintree ScientificRIS 100
Small animal lung inflation bulbHarvard Apparatus72-9083
SurgiSuite Multi-Functional Surgical Platform for Mice, with WarmingKent ScientificSURGI-M02Heated surgical platform
Test Leads 48" TEST LEAD BANANA - BlackMouser565-1440-48-0
Test Leads 48" TEST LEAD BANANA - RedMouser565-1440-48-2
Tracheal catheterΒ Exelint International2674622 G catheter
Wound closing system veterinary kitClay AdamsIN015Veterinary surgical stapling kit

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