Embed Size (px)
Citation preview
Determining the Oncological and Immunological Effects of Histotripsy for Tumor Ablation
Alissa D Hendricks-Wenger
Dissertation submitted to the faculty of the Virginia Polytechnic Institute and State University in partial fulfillment of the requirements for the degree of
Doctor of Philosophy In
Translational Biology, Medicine and Health
Eli Vlaisavljevich Irving Coy Allen
Eva Schmelz Scott Verbridge David Luyimbazi
April 26, 2021 Blacksburg, VA
Keywords: histotripsy, ablation, cancer, immunology, oncology models
This work is licensed under a Creative Commons Attribution 4.0 International
License by Alissa Danielle Hendricks-Wenger under http://creativecommons.org/licenses/by/4.0
DETERMINING THE ONCOLOGICAL AND IMMUNOLOGICAL EFFECTS OF
HISTOTRIPSY FOR TUMOR ABLATION
Alissa D Hendricks-Wenger
Abstract
Histotripsy is an emerging non-invasive, non-thermal, image-guided cancer
ablation modality that has recently been approved for its first clinical trial in the United
States (NCT04573881). Histotripsy utilizes focused ultrasound to generate acoustic
cavitation within a tumor to mechanically fractionate targeted tissues. While pre-clinical
work has demonstrated the feasibility of applying histotripsy to solid tumors including
primary liver and renal tumors, there is still a need to investigate the potential of
histotripsy to treat additional malignancies. In investigating the potential for treating
other malignancies there are two avenues that need to be considered: 1) the feasibility
for treating tissues with more complex stromal structures and 2) the ability of histotripsy
to modulate the tumor microenvironment. To determine the safety and feasibility of
additional applications of histotripsy, we conducted dose studies ex vivo on human
tumors and human liver to establish dosimetry metrics for applying histotripsy to more
fibrotic tumors such as cholangiocarcinoma while sparing nearby critical structures,
such as bile ducts and blood vessels. Learning the safety dose-margins from the
excised tissues, we performed an in vivo study using mice bearing patient-derived
xenograft cholangiocarcinoma tumors. With this model, we were able to demonstrate
our ability ablate the stiff cholangiocarcinoma tumors without causing any debilitating
off- target damage. To gain a more robust understanding of the effects of histotripsy
ablation on potentially difficult to treat tumors, we developed a porcine xenograft tumor
model and utilized veterinary cancer patients. These studies have helped established
protocols for utilizing histotripsy with ultrasound guidance to treat tumors that are more
difficult to treat and can withstand mechanical ablation, including pancreatic
adenocarcinoma, osteosarcomas, and soft tissue sarcomas. Pigs share many
similarities with human anatomy and physiology, making them an ideal model organism
for testing new medical devices and regimes for treating new targets. Using pigs, we
were able to establish a procedure to utilize histotripsy to target the pancreas in vivo
without causing any lasting or major side effects, such as off-target damage or
pancreatitis. One limitation to the porcine model and veterinary patients, is the limitation
of gaining rapid insight into the immunological effects of histotripsy. Established cancer
mouse models offer the opportunity to rapidly test many organisms with an intact
immune system. We used these mice to study pancreatic adenocarcinoma to determine
the immune response after histotripsy ablation. For these tumors the general response
was an increase in immune cell infiltration post-treatment and a shift in the tumor
microenvironment to a more anti-tumor environment. The results of this dissertation
provide insight into establishing protocols for treating new types of tumors with
histotripsy and immunological effects that lay groundwork for improving future co-
therapeutic treatment planning. Future work will aim to translate histotripsy into clinical
applications and determining co-therapies that can help control metastasis.
DETERMINING THE ONCOLOGICAL AND IMMUNOLOGICAL EFFECTS OF
HISTOTRIPSY FOR TUMOR ABLATION
Alissa D Hendricks-Wenger
General Audience Abstract
Histotripsy is a new medical therapy that can remove tumors without the need for
surgery, with the first clinical trial in the United States starting this year, 2021. This
therapy uses focused ultrasound waves to generate powerful microscopic bubbles that
can rapidly destroy targeted tissues with a high-degree of precision. Early studies on
histotripsy have demonstrated the ability of histotripsy to ablate tumors of the liver and
kidneys. In order to be able to fully use this therapy on more difficult to target and treat
cancers more studies are needed. Given that histotripsy uses physical forces to destroy
targets, stronger, more fibrotic tumors and cancers that have begun to spread
throughout the body will be more difficult to treat will need more than simple tumor
removal to better treat these patients. Therefore, when investigating new cancer
applications of histotripsy, it is important to consider the physical features of the tumors
as well as the ability of histotripsy to initiate an immune response against the cancer. To
determine the safety and feasibility of additional applications of histotripsy, we
conducted dose studies on excised human tumors and human liver to see what doses
of histotripsy are required to ablate stronger tumors, such as bile duct tumors. Learning
the potential safety margins of doses from the excised tissues, we conducted a study
using a mouse model to grow stiff, human tumors. With this model, we were able to
show that it is possible to ablate the stiffer tumors without causing any major off-target
damage. While it is useful to prove in excised tissues and mice that we can treat certain
tumors, there is an additional need to study the therapy in a model that is more similar
in size and anatomy to humans. Therefore, to gain a better understanding of the effects
of histotripsy on potentially difficult to target and ablate tumors, we developed a novel
porcine tumor model that can support the growth of human tumors and utilized
veterinary cancer patients. These studies have helped established protocols for utilizing
histotripsy to treat difficult to physically ablate tumors and difficult to ultrasound target
tumors, including pancreatic and bone cancers. Established cancer mouse models offer
the opportunity to rapidly test many organisms with an intact immune system. We used
these mice to study pancreatic cancer to determine the immune response after
histotripsy ablation. For this tumor type, while there were slight differences, the general
response was an increase in immune cell infiltration of the tumors post-treatment and a
shift to a stronger immune response against the tumor. The results of this dissertation
provide insight into establishing protocols for treating new types of tumors with
histotripsy and immune effects that lay groundwork for improving future co-therapeutic
planning. Future work will aim to translate histotripsy into clinical applications and
determining co-therapies that can help control body-wide disease.
vii
Dedication
To every person and dog that put up with my workaholic
personality, which allowed this body of work happen.
Thanks y’all.
viii
Acknowledgements
First and foremost, I would like to thank the first two who set me on the path that
is my PhD research. My thesis advisors Dr. Coy Allen and Dr. Eli Vlaisavljevich. Thanks
to Drs. Allen and Eli I have been able to accomplish a lot and grow as an independent
scientist. Unbeknownst to them (until reading this), I always joked that the only thing
that these two had in common was the PhDs at the end of their names, and this was my
saving grace for the past three years. While this caused me stress from time-to-time,
whenever I was stressed or frustrated, one of them always said what I needed (or
wanted) to hear.
I would also like to thank my current committee Dr. Eva Schmelz, Dr. David
Luyimbazi, and Dr. Scott Verbridge as well as Dr. Kenneth Oestrich, who was a
member of my committee until he moved to Ohio. I appreciate all the time that my
current and past committee members have set aside to listen to my research progress,
offer constructive critiques, and pose thought provoking questions.
Of course, I need to acknowledge the help and support that I have revied over
the years from the other students from both of my labs. I cannot image having survived
the past few years without everyone. It has really been like having two families to get
through the highs and lows of graduate school and real life. From the
engineering/TUSL/Eli/Vlaisavljevich lab (or whatever its being called now), thanks to
everyone who has helped with my projects or just been a person to chat with! Special
thanks to Alex Simon, who started working with histotripsy when I did, and whom I am
100% positive that without his help in the lab and out I would not have been able to get
ix
even half of this work done nor would it have been done with half the sanity. Thanks to
Lauren Arnold and Jessica Gannon who are obviously great engineers and both
inspirational, but also were never too busy to help improve all our days by complaining
about how busy we all were. I can also never forget to give a special shout out to Ruby
Hutchinson. While we never really worked together until she was walking out the door,
our ‘hikes’/glorified outdoor pandemic-safe walks and long conversations are
irreplaceable. From the Allen mucosal and cancer immunology lab, thanks to everyone
for helping with work as well as making work-life-balance a normal part of life. Special
thanks to Dr. Rebecca Brock, who started the TBMH program when I did and was an
impeccable roommate for 3 years. By joining the Allen lab first, Becca was the reason
that I met with Dr. Allen when looking to change labs and was therefore able to find my
graduate school niche. Additional thanks to Holly Morrison and Margaret Nagai-Singer
for always finding the positive outlook to difficult situations and being open to presenting
at yet another conference. Finally, of course, thanks to all the undergraduate and
medical students within both groups who have really been a crucial part of why I have
been able to complete this body of work. This especially goes out to Jackie Sereno,
Allison Zeher, Sofie Saunier, Jessica Gannon, Ruby Hutchison, and Pete Weber.
In addition to everyone within my labs, I also need to thank all our collaborators.
Thanks to the veterinary team, including Dr. Sheryl-Coutermarsh Ott, Dr. Joanne Touhy,
Dr. Nick Dervisis, and Dr. Shawna Klahn. Thanks to the various biomedical engineering
teams including Dr. Vincent Wang and his PhD student Chitra Meduri who have helped
with tissue mechanics work and Dr. Rafael Davalos and his PhD students Dr. Natalie
Beitel-White, Dr. Melvin Lorenzo, and Kenneth Aycock who assisted with the IRE
x
applications. Thanks to the clinicians from Carillion Clinic Dr. Douglas Grider and Dr.
David Luyimbazi, at Michigan Medicine Dr. Mishal Mendiratta-Lala, and at the
Universitat de Barcelona Dr. Joan Vidal-Jové.
I would also like to thank all of TBMH. Without Jay Read and Becky Langford I
would have been lost in the sea of paperwork. Also, thanks to Dr. Michael Friedlander,
Dr. Audra VanWort, Dr. Steve Poelzing, and Dr. Michelle Theus the directors of TBMH
during my time in the program. Additionally, I would like to thank all the students from
my cohort who helped me during our first-year courses and while preparing for our
qualifying exams.
Of course, an obligatory thanks to my husband Will and our dog Bunny, who put
up with some phases of neglect when I became a bit of a work-o-holic and still love me.
In general, thanks to all my friends and family who have been supportive and loving
throughout this whole process.
xi
Table of Contents
Abstract .......................................................................................................................... ii
General Audience Abstract ......................................................................................... iv
Dedication .................................................................................................................... vii
Acknowledgements .................................................................................................... viii
Table of Contents ......................................................................................................... xi
List of Figures ........................................................................................................... xvii
List of Tables ........................................................................................................... xxxii
List of Abbreviations .............................................................................................. xxxiii
Attributions for Co-Authored Papers ................................................................... xxxvi
Chapter 1. Introduction .............................................................................................. 1
1.1 Introduction .......................................................................................................... 2 1.2 Standard Ablation Procedures for Tumor Debulking and Immunomodulation .................................................................................................................................... 3 1.3 Histotripsy as a Tumor Ablation Therapy .......................................................... 5 1.4 Animal Models for Pre-Clinical Testing of Histotripsy ..................................... 7
1.5 Outline of this Dissertation ................................................................................. 9
1.6 References ......................................................................................................... 11
Chapter 2. Histotripsy for the Treatment of Cholangiocarcinoma Liver Tumors: In Vivo Feasibility and Ex Vivo Dosimetry Study ..................................................... 22
2.1 Abstract .............................................................................................................. 23 2.2 Keywords: .......................................................................................................... 24
2.3 Introduction ........................................................................................................ 25 2.4 Procedures ......................................................................................................... 27
2.4.1 Histotripsy Systems ............................................................................ 27 2.4.2 Pressure Calibration ........................................................................... 28
2.4.3 Histotripsy In Vivo CC Ablation Procedure ......................................... 29 2.4.4 Human Liver Tumor Specimens for Ex Vivo Treatments .................... 32
2.4.5 Histotripsy Ex Vivo Ablation Procedure .............................................. 33 2.4.6 Histology & Morphological Analysis .................................................... 36
2.5 Results ................................................................................................................ 37 2.5.1 Histotripsy Ablation of Subcutaneous PDX CC Tumors ..................... 37 2.5.2 Histotripsy Bubble Cloud Generation in Ex Vivo Tissue Specimens... 39
2.5.3 Histotripsy Ablation of Ex Vivo HCC Specimens ................................ 39 2.5.4 Histotripsy Ablation of Ex Vivo CLM Specimens ................................. 40
xii
2.5.5 Histotripsy Ablation of Ex Vivo CC Specimens ................................... 43 2.5.6 Histotripsy Ablation of Ex Vivo LP Specimens .................................... 44
2.6 Discussion ......................................................................................................... 46 2.7 Conclusion ......................................................................................................... 51 2.8 Acknowledgement ............................................................................................. 51 2.9 References ......................................................................................................... 53
Chapter 3. Histotripsy for Non-Invasive Ablation of Cholangiocarcinoma Carcinoma in a Patient-Derived Xenograft Model .................................................... 61
3.1 Abstract .............................................................................................................. 62 3.2 Keywords ........................................................................................................... 63 3.3 Introduction ........................................................................................................ 64 3.4 Materials and Methods ...................................................................................... 66
3.4.1 Tumor Injections and Health Monitoring ............................................. 66 3.4.2 Histotripsy Setup ................................................................................ 67
3.4.3 Histotripsy Treatment Procedures ...................................................... 68 3.4.4 Pilot Studies to Confirm Ablation Dose and Treatment Margins ......... 69
3.4.5 Survival Study ..................................................................................... 71 3.4.6 Necropsy and Histopathology ............................................................. 72
3.4.7 Statistics ............................................................................................. 73 3.5 Results ................................................................................................................ 74
3.5.1 Pilot Studies of Ablation Dose and Margins ........................................ 74
3.5.2 Survival Study ..................................................................................... 75 3.6 Discussion ......................................................................................................... 79
3.7 Conclusion ......................................................................................................... 82 3.8 Acknowledgment ............................................................................................... 83
3.9 References ......................................................................................................... 84
Chapter 4. Histotripsy Ablation in Preclinical Animal Models of Cancer and Spontaneous Tumors in Veterinary Patients ............................................................ 91
4.1 Abstract .............................................................................................................. 92 4.2 Introduction ........................................................................................................ 94 4.3 Procedures ....................................................................................................... 100
4.3.1 Histotripsy Systems and Pressure Calibrations ................................ 100 4.3.2 Hydrophone Focal Pressure Measurements, Beam Profiles, and Optical Imaging ......................................................................................... 103 4.3.3 Murine Tumor Models ....................................................................... 104 4.3.4 RAG2/IL2RG Deficient Porcine Tumor Model .................................. 106
4.3.5 Histotripsy Treatment of Porcine Pancreas and Liver ....................... 108 4.3.6 Spontaneous Tumors in Canine Patients ......................................... 110
4.3.7 Histotripsy Ablation of Spontaneous Canine Tumors ....................... 111 4.3.8 Histology & Morphological Analysis .................................................. 113
4.4 Results .............................................................................................................. 114 4.4.1 Histotripsy Systems for Small and Large Animal Studies ................. 114 4.4.2 Histotripsy Treatment of Subcutaneous Murine Tumors ................... 117
xiii
4.4.3 Characterization of RAG2/IL2RG Pig Tumor Model ......................... 120 4.4.4 Histotripsy Treatment of Porcine Pancreas and Liver ....................... 124
4.4.5 Veterinary Clinical Oncology Patient Populations for Histotripsy Development ............................................................................................. 125 4.4.6 Histotripsy Ablation of Spontaneous Canine Tumors ....................... 136
4.5 Discussion ....................................................................................................... 139 4.6 Conclusion ....................................................................................................... 149
4.7 Acknowledgment ............................................................................................. 150 4.8 References ....................................................................................................... 151
Chapter 5. Establishing a SCID-Like Porcine Model of Human Cancer for Novel Therapy Development with Pancreatic Adenocarcinoma and Irreversible Electroporation .......................................................................................................... 169
5.1 Abstract ............................................................................................................ 170 5.2 Keywords ......................................................................................................... 171
5.3 Introduction ...................................................................................................... 172 5.4 Materials And Methods ................................................................................... 176
5.4.1 Generation of Immunodeficient Pigs ................................................. 176 5.4.2 Animal Care ...................................................................................... 178
5.4.3 Tumor Injection and Monitoring ........................................................ 178 5.4.4 Immunohistochemistry ...................................................................... 179 5.4.5 Ultrasound Imaging .......................................................................... 180
5.4.6 Tissue Handling and Electroporation ................................................ 180 5.5 Results .............................................................................................................. 183
5.5.1 Generation of immunodeficient pigs carrying targeted deletions in RAG2/IL2RG using the CRISPR/Cas9 system .......................................... 183
5.5.2 Panc01 tumors successfully engrafted in all pigs carrying the targeted RAG2/IL2RG deletion ................................................................................ 185
5.5.3 Panc01 tumors are histopathologically similar to Pan02 and PDX tumors generated in mice .......................................................................... 187 5.5.4 Panc01 tumors demonstrate similar electrical properties to primary patient derived pancreatic cancer tissue and healthy porcine tissues ....... 191
5.6 Discussion ....................................................................................................... 195 5.7 Abbreviations ................................................................................................... 199 5.8 Funding Sources ............................................................................................. 199 5.9 References ....................................................................................................... 200 5.10 Supplemental Figure ..................................................................................... 209
Chapter 6. Feasibility of Histotripsy Pancreas Ablation in a Large Pig Model . 210 6.1 Abstracts .......................................................................................................... 211
6.2 Keywords: ........................................................................................................ 212 6.3 Introduction ...................................................................................................... 213 6.4 Materials and Methods .................................................................................... 216
6.4.1 Animal Monitoring and Anesthesia ................................................... 216 6.4.2 Histotripsy Systems and Pressure Calibrations ................................ 218
xiv
6.4.3 Hydrophone Focal Pressure Measurements and Beam Profiles ...... 219 6.4.4 Histotripsy In Vivo Pancreas Ablation Procedure ............................. 220
6.4.5 Computed Tomography Imaging ...................................................... 222 6.4.6 Necropsy & Histological Analysis ..................................................... 223
6.5 Results .............................................................................................................. 223 6.5.1 Establishing Safe, Non-Invasive Therapeutic Protocol ..................... 223 6.5.2 Physiological Outcomes ................................................................... 226
6.6 Discussions ..................................................................................................... 230 6.7 Conclusion ....................................................................................................... 232 6.8 Acknowledgements ......................................................................................... 232 6.9 References ....................................................................................................... 233
Chapter 7. Immunological Effects of Histotripsy for Cancer Therapy ............... 239
7.1 Abstract ............................................................................................................ 240 7.2 Keywords: ........................................................................................................ 241
7.3 Introduction ...................................................................................................... 242 7.4 The Role of Ablation in Immunomodulation ................................................. 243
7.4.1 The Tumor Microenvironment and Tumor Associated Immune Cells 247 7.4.2 Mechanisms for Shifting the Tumor Microenvironment to Pro-Inflammatory .............................................................................................. 252
7.5 Histotripsy: Non-Thermal, Non-Invasive Tumor Ablation ............................ 255 7.6 Immunologic Effects of Histotripsy ............................................................... 258
7.6.1 Decreases in Pro-Tumor Immune Cells ............................................ 258 7.6.2 Histotripsy Produces Damage Associated Molecular Patterns that Directly Increase Local Cellular Immune Responses ................................ 261 7.6.3 Pro-Inflammatory Cytokines and Chemokines Associated with Histotripsy Based Tumor Ablation Modalities ............................................ 264 7.6.4 Histotripsy Significantly Alters Immune Cell Populations Systemically and in the Tumor Microenvironment .......................................................... 267 7.6.5 Improved Engagement of the Adaptive Immune System and Increases in the Systemic Anti-Tumor Immune Response ........................................ 268
7.6.6 Combination Therapy with Checkpoint-Inhibitors ............................. 274
7.7 Conclusion and Future Outlook ..................................................................... 277 7.8 Acknowledgements ......................................................................................... 278 7.9 References ....................................................................................................... 279
Chapter 8. Histotripsy Ablation Alters the Tumor Microenvironment and Promotes Immune System Activation in a Subcutaneous Model of Pancreatic Cancer 299
8.1 Abstract ............................................................................................................ 300
8.2 Keywords ......................................................................................................... 301 8.3 Introduction ...................................................................................................... 302 8.4 Procedures ....................................................................................................... 304
8.4.1 Tumor Injections and Monitoring ...................................................... 304 8.4.2 Histotripsy Set Up ............................................................................. 305
xv
8.4.3 n Vitro Ablations Treatment Parameters ........................................... 307 8.4.4 In Vivo Histotripsy Treatment ........................................................... 309
8.4.5 Determining DNA Release and Quality ............................................ 312 8.4.6 Determining Protein Release and Quality ......................................... 312 8.4.7 Histopathology .................................................................................. 312 8.4.8 Profiling Gene Expression and Pathway Analysis ............................ 313 8.4.9 Flow Cytometry Panel Staining......................................................... 314
8.4.10 HMGB1 Serum ELISA .................................................................... 315 8.4.11 Statistics ......................................................................................... 315
8.5 Results .............................................................................................................. 316 8.5.1 Different Ablation Modalities Show Differential Release of Damage Associated Molecular Patterns (DAMPs) and Potential Antigens .............. 316
8.5.2 Histotripsy is an Effective Tumor Ablation Modality in the Subcutaneous Pan02 Model ..................................................................... 318
8.5.3 Histotripsy Ablation Results in Increased Acute Cellular Immune Response .................................................................................................. 321
8.5.4 Histotripsy Alters Immune Cell Composition in the Tumor Microenvironment ...................................................................................... 323
8.6 Discussion ....................................................................................................... 325 8.7 Conclusion ....................................................................................................... 331 8.8 Acknowledgment ............................................................................................. 332
8.9 References ....................................................................................................... 333 8.10 Supplemental Table and Figure Legends .................................................... 342
Chapter 9. Conclusion and Future Directions ..................................................... 344 9.1 Introduction ...................................................................................................... 345
9.2 How can we safely and efficacious ablate more difficult tumors? .............. 345 9.3 Can we use a large animal model to develop histotripsy for acoustically difficult to target tumor locations? ...................................................................... 346 9.4 How do different tumor types immunologically respond to histotripsy? ... 348 9.5 Overall Conclusion .......................................................................................... 350
Appendix A. Employing Novel Porcine Models of Subcutaneous Pancreatic Cancer to Evaluate Oncological Therapies ............................................................. 351
A.1 Abstract ........................................................................................................... 352 A.2 Keywords ......................................................................................................... 352 A.3 Introduction ..................................................................................................... 353 A.4 Materials .......................................................................................................... 355
A.4.1 Generating RAG2/IL2RG Deficient Pigs .......................................... 355 A.4.2 Maintaining Cells in Culture ............................................................. 356
A.4.3 Harvesting Cells for Injection............................................................ 357 A.4.4 Sterile Passing in of the Cells........................................................... 357 A.4.5 Injecting Tumors ............................................................................... 357 A.4.6 Monitoring Tumors and Health ......................................................... 357 A.4.7 Ultrasound Imaging (see Note 6) ..................................................... 358
xvi
A.4.8 Necropsy .......................................................................................... 358 A.5 Methods ........................................................................................................... 359
A.5.1 Generating RAG2/IL2RG Deficient Pigs .......................................... 359 A.5.2 Maintaining Cells in Culture ............................................................. 360 A.5.3 Harvesting Cells for Injection............................................................ 360 A.5.4 Sterile Passing in of the Cells........................................................... 361 A.5.5 Injecting Tumors ............................................................................... 362
A.5.6 Monitoring Tumors and Health ......................................................... 363 A.5.7 Ultrasound Imaging .......................................................................... 364 A.5.8 Necropsy .......................................................................................... 365
A.6 Notes ................................................................................................................ 366 A.7 References ....................................................................................................... 369
xvii
List of Figures
Figure 1-1 Histotripsy is a non-invasive ablation method that uses focused ultrasound to
generate a cavitation bubble cloud, resulting in the complete destruction of targeted
tissues with high precision. .............................................................................................. 5
Figure 2-1 In Vivo Ablation Setup. (A) Histotripsy murine experiments were conducted
using a custom 1MHz small animal histotripsy system. (B) Subject’s tumors were
submerged in degassed water at the transducer focus in order for (C) histotripsy to be
applied to the tumor non-invasively guided by real-time ultrasound imaging. ............... 30
Figure 2-2 Ex Vivo Tissue Ablation Setup. Histotripsy dose experiments were conducted
using a 700kHz transducer with the focus aligned in the center of the targeted tissue,
with guidance from ultrasound imaging aligned orthogonally (A-B). Tissues were
imbedded within the center of a gelatin phantom (C). ................................................... 32
Figure 2-3 Dotted orange circles on US images (A-D) indicate approximate region of
ablation. Low magnification H&E images shows ablation margins (E-H). High
magnification H&E images of control and ablated region (J-L). Black boxes on 10x
images indicate area magnified in respective 40x images. Scale bar on 10x H&E
images = 100um (E-H), 40x scale bar = 50um (I-J). ..................................................... 38
Figure 2-4 Comparison of untreated human HCC tumors with ex vivo ablations at 500
pulses/point, 1000 pulses/point, and 2000 pulses/point. 10x magnification scale bar=
100um (A-D), 40x scale bar= 50um (E-H). Black boxes on 10x images indicate area
magnified in respective 40x images. Histology images are representative of the samples
treated. .......................................................................................................................... 40
xviii
Figure 2-5 Comparison of untreated human CLM tumors with those treated at 500
pulses/point, 1000 pulses/point, 1500 pulses/point, and 2000 pulses/point. 10x
magnification scale bar= 100um (A-E), 40x scale bar= 50um (F-J). Black boxes on 10x
images indicate area magnified in respective 40x image. Histology images are
representative of the samples treated. .......................................................................... 41
Figure 2-6 Comparison of untreated human CC tumors with those treated at 1000
pulses/point, 1500 pulses/point, 2000 pulses/point, and 4000 pulses/point. 10x
magnification scale bar= 100um (A-E), 40x scale bar= 50um (F-J). Black boxes on 10x
images indicate area magnified in respective 40x image. Histology images are
representative of the samples treated. .......................................................................... 42
Figure 2-7 Supplemental Figure S1. Single sample of Cholangiocarcinoma Ablated at
4000 pulses/point illustrating fibrosis dependence to susceptibility. Region of fibrosis (A)
showed no damage, while less fibrotic region (B) had sporadic ablated foci. High
magnification of the region in B indicated by the black box is Figure 7E 4000
pulses/point location. Scale bars = 500um. * Indicate ablated region. .......................... 44
Figure 2-8 Non-steatotic liver parenchyma treatment at 250 pulses/point, 500
pulses/point, and 1000 pulses/point demonstrating less uniform ablations. 10x
magnification scale bar= 100um (A-D), 40x scale bar= 50um (E-H). Black boxes on 10x
images indicate area magnified in respective 40x images. Histology images are
representative of the samples treated. .......................................................................... 45
Figure 2-9 Steatotic liver parenchyma treatment at 250 pulses/point, 500 pulses/point,
and 1000 pulses/point demonstrating less uniform ablations. 10x magnification scale
bar= 100um (A-D), 40x scale bar= 50um (E-H). Black boxes on 10x images indicate
xix
area magnified in respective 40x images. Arrows indicating example cells within an
ablated region that appear to be intact. Histology images are representative of the
samples treated. ............................................................................................................ 46
Figure 3-1 Rodent system treatment set up (A). Ultrasound imaging confirmed presence
of bubble cloud (indicated by the orange arrow) within CC tumors during treatment (B).
For treatment automation, each tumor was defined as a spheroid divided into disks (C).
Each treatment disk consisted of a series of points that were used for raster scanning
(D). White arrow in (D) represents the direction of therapeutic ultrasound wave
propagation. .................................................................................................................. 70
Figure 3-2 Schematic showing the planned ablation volumes treatments utilized for this
study. Negative-margin treatments were set to have the entire treatment contained
within the tumor; zero-margin treatments were set to treat as close to 100% of the tumor
as possible without including a significant amount of non-tumor tissue; and positive-
margin treatments were set to treat the entire tumor along with sufficient margin that
encompassed enough non-tumor tissue to ensure complete ablation of the entire tumor.
Arrows indicate tumor tissue not targeted by each planned ablation zone while * indicate
intestinal tissue within the treatment margin. Figure created using Biorender.com. ...... 72
Figure 3-3 Histological examination revealed undisturbed tumor stroma in acute
untreated mice (A) and completely acellular targeted regions in treated tumors (B).
Comparing to before treatment (C), the hypoechoic regions were identifiable in the
ultrasound imaging after treatment (D). The treatment area, represented by the dotted
yellow circle, closely matched the negative-margin planned ablation volume on
ultrasound imaging. ....................................................................................................... 73
xx
Figure 3-4 Using positive, more aggressive margins (n=2) there was more significant
skin lesions and subcutaneous bruising that extended well beyond the treatment area
across the abdomen (A). Darkened, parrelel lesions that correspond to treatment disks
are emphasized with purple arrows. At necropsy, these more aggressive treatments
revealed free fluid and intestinal contents within the abdomen and a clear intestinal
perforation, indicated by the green arrow (B). ............................................................... 74
Figure 3-5 No significant skin breaks were observed during treatment (A). Immediately
after treatment, there was slight bruising surrounding the tumors, indicated by the blue
arrow (B). Three days after treatment there was no skin damage in the untreated mice
(C) and small, pitted scabs over the treated areas, noted by the red arrow (D). ........... 76
Figure 3-6 Average tumor diameter (mean + SEM) reduction after treatment, after
treatment the difference between groups was significant at every time point (* p<0.05)
(A). Individual treated and untreated mouse tumor measurements, emphasizing that
multiple treated animals reached a tumor diameter of zero (B). Vertical dotted line
emphasizes treatment on day 8. Horizontal dotted line at 14mm emphasizes maximum
tumor diameter that any individual animal was allowed to reach in this study based on
our IACUC protocols. .................................................................................................... 77
Figure 3-7 Progression free and overall survival increased (p=0.09 and 0.03
respectively) in treatment group compared to untreated. Progression free survival was
defined as surpassing 120% of the average tumor diameter the day of treatment (5.29
mm). Vertical dotted line emphasizes treatment on day 8. ............................................ 78
Figure 3-8 Histological examination revealed undisturbed tumor stroma in chronic (A)
untreated mice and tumor regrowth in chronic treated tumors (B). The liver (C-D) and
xxi
lung (E-F) parenchyma in both untreated (E,C) and treated (D,F) chronic mice were
found to be benign. Scale bar = 500um. ....................................................................... 79
Figure 4-1 Example Small Animal Histotripsy System. (A) A small animal system
consisting of a 1) 1 MHz therapy transducer with an 2) imaging transducer coaxially
aligned and mounted to a 3) 3D motorized positioning system to deliver hisotripsy to
target submerged in degassed water on a 5) subject stage. A custom-built electronic
driving system is controlled with computer software. (B) Linear ultrasound imaging
probe coaxially aligned with 1 MHz therapy transducer used to (C) treat tumors in-vivo.
(D) Acoustic waveform for 1 MHz transducer collected with a hydrophone at a peak
negative pressure of ~10 MPa. (E) 1 MHz bubble cloud captured on a high-speed
camera. ....................................................................................................................... 101
Figure 4-2 Example Large Animal Histotripsy System. (A) Histotripsy system designed
for liver cancer ablation consisting of 1) therapy and 2) imaging transducers attached to
an 3) articulating arm with a 4) motorized micro-positioner controlled by 5) custom
software. (B) Image of therapy transducer with co-axially aligned imaging probe used for
(C) previous in vivo liver studies. (D) Acoustic waveform for 500 kHz transducer
collected with a hydrophone at a peak negative pressure of ~10 MPa. (E) 500 kHz
bubble captured on a high-speed camera. .................................................................. 102
Figure 4-3 Histotripsy Tumor Ablation in Murine Models. (A) Subcutaneous murine
pancreatic Pan02 tumor. (B) Representative primary human pancreatic tumor. (C)
Human CC PDX murine xenografted tumor. (D) Ultrasound images of histotripsy bubble
clouds forming in RCC, (E) 4T1, and (F) CC PDX tumors. (G-I) Histology demonstrating
full ablation within histotripsy targeted (G) RCC, (H) 4T1, and (I) CC PDX tumors. .... 118
xxii
Figure 4-4 RAG2/IL2RG Immunocompromised Pig Model for Cancer Research. (A)
Example of a Pan01 (pancreatic cancer) subcutaneous tumor nodule growing behind
the ear and (B) once excised. (C) Control Matrigel plugs exhibits no growth. (D, G) U-
251 Human Glioblastoma, (E, H) 4T1 murine breast cancer, and (F, I) HepG2 human
HCC are shown by histology under low and high magnification, respectively. ............ 121
Figure 4-5 Histotripsy Treatment of Porcine Pancreas and Liver. (A) Image of the water
bowl and site preparation with fluid repellent adhesive surgical drape for liquid
containment during histotripsy. (B) Acoustic window for pancreas ultrasound treatment
identified on pre-treatment imaging. (C) Gas in the stomach and GI tract encountered
prior to histotripsy treatment. (D-E) Guided by real-time ultrasound imaging, histotripsy
generated a clearly defined bubble cloud in (D) liver and (E) pancreas. (F) Gross image
of 0.5 cm lesion in pancreas following histotripsy. (G) Off-target effects around the
stomach and GI tract. (H-I) Representative histopathology images confirm regions of
ablation in the pancreas. ............................................................................................. 123
Figure 4-6 Example Spontaneous Tumors in Canines. (A) Soft Tissue Sarcoma (STS)
presented as a firm, fixed, painful soft tissue mass at the right proximal forelimb of a
dog. (B) Contrast-enhanced CT consistent with a soft tissue attenuating and moderately
contrast enhancing mass (yellow asterisk). The mass lies lateral to the triceps muscle
and arises from the ascending pectoral muscle. (C) HIFU treatment of the tumor to
partial ablation. An Echopulse unit by Theraclion was used. (D) Gross appearance of a
cross-section of the tumor, 4 days post HIFU treatment. The yellow arrows point to the
ablated tumor volume. (E) CT image of a rostral maxillary tumor in a dog, indicated in
circled area. Note the aggressive invasion of the tumor into the maxilla, causing bone
xxiii
lysis. (F) CT image of a humeral osteosarcoma in a dog, indicated by the arrow. Note
the aggressive nature of the tumor, causing bone lysis and proliferation. (G) CT image
of a liver tumor in a dog, indicated by the circle. (H) CT image of a pancreatic tumor in a
dog, indicated by the arrow. (I) CT image of a metastatic lymph node in the same
patient, indicated by a circle. ....................................................................................... 130
Figure 4-7 Histotripsy Treatment of Canine Soft Tissue Sarcoma. (A-B) Image of
spontaneous soft tissue sarcoma with estimated tumor size of ~7-8 cm in diameter. (C)
Contrast-enhanced CT collected before histotripsy treatment. (D) Histotripsy treatment
experimental set-up. (E) Real-time ultrasound image collected during histotripsy
treatment showing the bubble cloud visible as a hyperechoic, oscillating zone (arrow).
(F) Contrast-enhanced CT collected 4 days after histotripsy treatment with decreased
contrast uptake visible in the zone of ablation. (G) Gross morphological analysis of
resected canine soft tissue sarcoma. Box indicates treatment region, identified by
location and presence of tissue necrosis and hemorrhage. (H) H&E image of untreated
canine soft tissue sarcoma (20x magnification) with intact cells and undisturbed
extracellular matrix is observable. (I) H&E image of canine soft tissue sarcoma tissue
treated with histotripsy (40x magnification) showing extensive necrosis and hemorrhagic
appearance. ................................................................................................................ 136
Figure 4-8 Histotripsy Treatment of Canine Osteosarcoma. (A-B) Image of an
undisturbed spontaneous canine osteosarcoma in the left hind limb. Significant soft
tissue swelling was present, creating an estimated zone of ~30 cm of affected tissue.
(C) Contrast-enhanced CT collected before histotripsy treatment. (D) Histotripsy
treatment experimental set-up. (E) Real-time ultrasound image collected during
xxiv
histotripsy treatment with the bubble cloud visible as a hyperechoic, oscillating zone
(arrow). (F) Contrast-enhanced CT collected 1 day after histotripsy treatment. .......... 137
Figure 5-1 Generation of RAG/ IL2RG knockout pigs. (A) Schematic approach for
RAG2/IL2RG knockout pig production. First, the sgRNA and Cas9 mRNA was
synthesized via in vitro transcription. Second, an optimal concentration of sgRNA and
Cas9 was injected into presumable zygotes. Third, the injected embryos were
transferred into a surrogate gilt. At the end of gestation, piglets were born via
hysterectomy and maintained in gnotobiotic isolators. (B) Summary of piglet genotypes
in IL2RG and RAG2 gene. Six piglets were born and analyzed. During PCR analysis,
RAG2 was not amplified from the Piglet #2 DNA sample, suggesting a large deletion.
Biallelic genotype indicates two differently modified alleles. Homozygous indicates one
type of modified alleles. (C) Genotype of Piglet 1 as a representation of small deletions
or insertions introduced by the CRISPR/Cas9 system. ............................................... 182
Figure 5-2 Confirmation of SCID-Like Phenotype. FACS analysis to detect the presence
of lymphocytes in RAG2/IL2RG knockout piglets. Compared to wild-type control, SCID
piglet possessed lower level of B (CD79A), T (CD3E), and NK (CD3E-/CD56+) cells. 185
Figure 5-3 Successful Engraftment of Panc01 Cells. (A) Tumor growth was measured
three times per week for each pig. All animals developed palpable tumors that
continued to grow over the course of the study. (B) Pig weight was monitored
throughout the duration of the study and utilized as a surrogate for animal morbidity. 185
Figure 5-4 Subcutaneous Tumors Generated Ample, High-Quality Pancreatic Cancer
Tissue for Subsequent Ex Vivo Tumor Ablation Assessments. (A) Tumor growth was
readily observable above the skin behind the ear where the Panc01 cells were injected.
xxv
(B) No observable foci were located in regions where Matrigel alone (control) was
injected. (C) Ultrasound image confirmed the volumetric growth of the tumors. The
tumor is outlined here in white for clarity. (D-E) Necropsy of the tumors confirmed that
(D) they were confined to the dermis and (E) approximately the same size as measured
extra-dermally. ............................................................................................................ 187
Figure 5-5 Histopathology Comparison of Pre-Clinical Pancreatic Cancer Animal
Models. Human PDX, mouse Pan02, and human Panc01 were compared. (A)
Subcutaneous tumors in the mouse Pan02 model are densely cellular often with central
areas of necrosis (asterisk). Individual cells are spindle-shaped and arranged in vague,
irregular streams. (B) Subcutaneous tumors in the PDX mice are also highly cellular and
also exhibit foci of necrosis (asterisk). At higher magnification, these cells are more
polygonal and are often arranged to form irregular tubular structures with lumens
containing inflammatory cells and debris (arrows). (C) Subcutaneous tumors in the
porcine Panc01 model are also densely cellular and variably exhibit foci of necrosis.
Individual cells are more polygonal in shape and rarely attempt to recapitulate glandular
structures (arrows). ..................................................................................................... 188
Figure 5-6 Immunohistochemical Confirmation of Ductal Cells in Pre-Clinical Pancreatic
Cancer Animal Models. (A) Human PDX, (B) mouse Pan02, and (C) human Panc01
were compared for CK19 expression. Brown color indicates positive color. Nuclei are
stained in dark blue, and remaining cell features are highlighted in light blue. ............ 189
Figure 5-7 Primary human, SCID-like Panc01, and murine PDX tumors demonstrate
similar conductivity increases following high voltage pulsed electric field. Ex vivo tissue
samples directly from patients, RAG2/IL2RG animals, or NSG mice were sectioned and
xxvi
fit within a PDMS mold to impose a cylindrical shape factor. After sample preparation,
individual samples were exposed to IRE pulses with varying electric field magnitudes.
A) Summary of initial conductivities calculated from normal and cancerous SCID-like
(black), human (red), and murine PDX (gray) tissue. B) The conductivity for each tissue
type increased with the applied electric field. C) Percent increase in tissue conductivity
at varying fields was determined from the sample-specific pre-pulse values. D) The
adjusted conductivity is calculated from the percent difference values; here, the
conductivities are normalized to the average initial tissue conductivity and adjusted
based on the percent change. ..................................................................................... 191
Figure 5-8 Impedance for healthy tissue excised from the RAG2/IL2RG pigs correlate
adult swine and humans. A Gamry Reference 600 potentiostat (Gamry, Warminster,
PA, US) was used to record tissue impedance within a frequency range of 103 to 106
Hz. The real part of the tissue impedance was used in the calculation of tissue
conductivity for further comparison to values previously reported in literature. A)
Comparison of electrical conductivity from SCID-like pig tissue to healthy porcine
(pancreas/brain) or human (prostate) tissue. The B) pancreatic, C) brain, and D)
prostate tissue demonstrate similar electrical behavior to their normal counterparts
across a wide frequency range. Each data point (dot) represents reported values for
normal tissue conductivity, while the black line represents data gathered in this study
and the red line represents normal porcine tissue. (*p < 0.05). ................................... 192
Figure 5-9 Supplemental Figure 5.S1. Genotyping of piglets carrying modified IL2RG
and RAG2 gene. Six piglets were born and analyzed. During PCR analysis, RAG2 was
xxvii
not amplified from the Piglet #2 DNA sample, suggesting a large deletion. No wild-type
allele was identified. .................................................................................................... 209
Figure 6-1Study timeline demonstrating the relative time for treatment, custard feeding,
blood draws, and CT imaging. ..................................................................................... 217
Figure 6-2 HistoSonics cart with 500 kHz transducer and coaxial US imaging probe
mounted onto a micropositioner (A). Freehand imaging by radiologist to identify acoustic
window (B). UMC bowl with therapy transducer submerged via adjustable arm (C). .. 218
Figure 6-3 Bubble cloud formed in liver on US imaging from HistoSonics system (pig 1,
8-6)(A). No bubble cloud visualized on US imaging and did not clearly identify the
pancreas without pudding (pig 6, 8-13)(B). Bubble cloud generated inside pancreas
where pig was fed pudding prior to histotripsy treatment (28-53) (C). ......................... 224
Figure 6-4 Intestinal gas interference with treatment prevented ideal imaging quality and
bubble formation during treatment leading to pre-focal intestinal bruising (A-B) more
highly concentrated in the more gaseous large intestine, indicated by the green arrow,
and near the targeted region, indicated by the blue arrows. Post treatment H&E
histopathology indicated that from a sample of the bruised colon, the muscularis was
not damaged but there was a large amount of ablation within the submucosa (C-D). 225
Figure 6-5 US imaging of pancreas before histotripsy (A) and after treatment with clear
hypoechoic region identifying the targeted ablated volume (B). Dotted red circle
indicated the approximate region treated .................................................................... 226
Figure 6-6 With laxative, simethicone, and custard pretreatment protocol, there was
signifcatly less bruising in preforcal bowel (A) indicated by the green arrow. Bruising
xxviii
around the treatment area was more concentrated than on the bowel, but was still less
intense than the acute group (B), bruising indicated by blue arrow. ............................ 227
Figure 6-7 CT taken in the dorsal recumbent position (A) one week post treatment
shows no free fluid in the peritoneal cavity (B) ............................................................ 228
Figure 6-8 White blood cell, amylase, and lipase serum levels for 1-week survival
animals. ....................................................................................................................... 228
Figure 6-9 1-week post treatment there was no signs of morbidity within the pancreatic
tissue. Images representative of two animals. ............................................................. 229
Figure 6-10 Necropsy reviled no gross damage within the abdomen in situ (A-B). When
excised in tota no damage was found on the exterior of the liver (C-D), the spleen (E),
nor the pancreas (F-G). ............................................................................................... 229
Figure 7-1 Focal Treatment of Targeted Tumor and Mechanism for Systemic Tumor
Control. The right diagram demonstrates the principal of the abscopal effect, where the
treatment of one tumor can cause other tumors in the body to shrink or be eliminated to
varying degrees due to immune system engagement. The left diagram depicts the
simplified mechanism for achieving a systemic effect from focal therapies. DC: Dendritic
cell. MHC: Major Histocompatibility Complex. TCR: T cell receptor. ........................... 245
Figure 7-2 Pro-Tumor Immune Cells. CD66, CD66b, and CD15 are human specific
markers. Gr-1 represents both Ly6C and Ly6G, therefore does not distinguish the
MDSC subtypes, MDSCs are monocytic when Ly6C+ and granulocytic when Ly6G+.
Most of the markers available for pro-tumor immune cells are surface receptors and
ligands, exceptions listed here include the intracellular protein Arg1 and the transcription
factor FOXP3. ............................................................................................................. 249
xxix
Figure 7-3 The Role of Check Point Inhibitors. Antigen Presenting Cells (APC). ........ 253
Figure 7-4 Histotripsy Schematic. Therapeutic ultrasound transducer positioned outside
of the body, focuses ultrasound waves within a targeted tissue generating a cavitation
bubble cloud. ............................................................................................................... 256
Figure 7-5 Schematic of Histotripsy’s Postulated Roles in Immune System Modulation.
Immune processes, cells, and molecules that have been shown to be modulated by
focused ultrasound therapies after mechanical ablation are summarized based upon
immune system functions: pro-tumor immune cells, cellular immunity, and systemic
immunity. ..................................................................................................................... 259
Figure 8-1 In Vivo Experimental Setup. Murine histotripsy experiments were conducted
using a 1Mhz transducer (A). Timing for treatment, euthanasia, and data collection were
set as diagramed (B). Days with histology collection are noted by cassettes, flash frozen
tumors noted by tubes with tumors, serum noted by tubes with serum, and flow
cytometry is noted by flow charts. Tumors located with ultrasound imaging were used
for planning automated ablation disks (C) and raster scan plots (D). Therapy was
guided by co-axially aligned ultrasound imaging (E). Red arrow indicates a bubble cloud
that was generated during treatment. .......................................................................... 306
Figure 8-2 Treatment flow and dosages for various ablation modalities (A). Histotripsy
experiments on cell suspensions were conducted using a 1Mhz transducer (B). ....... 308
Figure 8-3 Cell suspension ablations resulted in partial and full ablations (A). DNA
release was quantified (B) and visualized (C). Protein release was quantified (D) and
the relative release of the antigen HA was quantified from western blot bands (E).
Lowercase letters on top of bars indicate significance; bars with the sample letter
xxx
designation are not significant while those that do not share a letter are significant
(p<0.05). ...................................................................................................................... 316
Figure 8-4 Ultrasound images of tumor before (A) and after (B) treatment, exhibiting
more hypoechoic central region post-histotripsy. Orange shapes on ultrasound indicate
location of tumors. Green circles indicate location of bubble cloud determined in water
prior to treatment, and were utilized for treatment planning. Histology to tumors without
(C) and with (D) treatment show decreased cellular detail after treatment. Dotted black
line outlines the necrotic core of the tissue. Scale bar on H&E images = 500 um. ...... 319
Figure 8-5 After treatment of tumors on day 8, as indicated by the black arrow, the
reduction of tumor volume was indicated by caliper measurements (A). Changes health
observed in tumor-progression free survival (B) and general survival (C). .................. 320
Figure 8-6 Analysis of mRNA expression in treated tumors showed regulation of
immune pathways (A). Regulated pathways interact with each other and are up
regulated in the acute group and downregulated in the chronic group (B). Serum
HMBG-1 levels (C) correlate with the mRNA upregulation of HMGB-1 associated
pathways. .................................................................................................................... 321
Figure 8-7 Single cell suspension from treated and untreated tumors were collected at
24 hours, 7 days, and 14 days after treatment and were stained as described for flow
cytometry to identify innate immune cells. Percentage of each innate immune cell
population analyzed was calculated as part of the total CD45+ cells stained. Example
flow cytometry plots from treated and untreated tumors at 14 days after treatment. ... 324
Figure 8-8 Single cell suspension from treated and untreated tumors were collected at
24 hours, 7 days, and 14 days after treatment and were stained as described for flow
xxxi
cytometry to identify adaptive immune cells. Percentage of each adaptive immune cell
population analyzed was calculated as part of the total viable cells stained (gated as
singlets). The ratio of CD4+/CD8+ T cells was calculated as a simple ratio. Example
flow cytometry plots from treated and untreated tumors at 14 days after treatment. ... 325
Figure 8-9 Supplemental Figure 1: Whole western blot images analyzing HA antigen
released in ablated cell supernatants. ......................................................................... 342
Figure 0-1: Creating “Tent” to Ensure Subcutaneous Injection. By pulling the pig’s ear
forward, a small tent of skin will pull up forming a small tent. To ensure a subcutaneous
injection, have the bevel of the needle enter “tent” following a path indicated ............. 362
Figure 0-2: Measuring Subcutaneous Tumor Growth in Gnotobiotic Porcine Isolators.
(A) Subcutaneous tumors (indicated by arrow) grown behind the ear can be easily
viewed within two days post injection and (B) measured with plastic Vernier calipers. (C)
The .............................................................................................................................. 363
xxxii
List of Tables
Table 2-1 Ex Vivo sample numbers by tissue type and Dose --------------------------------- 36
Table 5-1 Target sequences of RAG2 and IL2RG sgRNAs. ---------------------------------- 176
Table 5-2 Primers used to amplify sgRNA target regions. ------------------------------------ 178
Table 8-1 Subject Numbers Per Experimental Group ------------------------------------------ 309
Table 8-2 Flow Cytometry Immune Cell Markers ------------------------------------------------ 313
Table 8-3 Supplemental Table 1: Genes analyzed from SuperArry rtPCR and calculated
fold regulation. -------------------------------------------------------------------------------------------- 343
xxxiii
List of Abbreviations
ACCRC: Animal Cancer Clinic & Research Center
ANOVA: Analysis of Variance
APC: Antigen Presenting Cell
ATCC: American Type Culture Collection
BH: Boiling Histotripsy
CBC: Complete Blood Count
CC: Cholangiocarcinoma
CCH: Cavitation Cloud Histotripsy
CLM: Colorectal Liver Metastasis
COTC: Comparative Oncology Trials Consortium
CRISPR: Clusters of Regularly Interspaced Short Palindromic Repeats
CT: Computed Tomography (imaging)
DAMP: Damage Associated Molecular Pattern
DC: Dendritic Cell
DNA: Deoxyribose Nucleic Acid
ERF: Epidermal Growth Factor
FOPH: Fiber Optic Probe Hydrophone
FPGA: Field Programmable Gate Array
FWHM: Full width Half Maximum
H&E: Hematoxylin and Eosin
HA: Hemagglutinin
HCC: Hepatocellular Carcinoma
xxxiv
HIFU: High Intensity Focused Ultrasound
IACUC: Institutional Animal Care and Use Committee
IL: Interleukin
IRB: Institutional Review Board
IRE: Irreversible Electroporation
MDSC: Myeloid Derived Suppressor Cell
MHC: Major Histocompatibility Complex
mHIFU: Mechanical High Intensity Focused Ultrasound
MRI: Magnetitic Resonance Imaging
NCI: National Cancer Institute
NK: Natural Killer (cell)
NSG: Nod-SCID Gamma
OCM: Oncopig Cancer Model
OS: Osteosarcoma
PDX: Patient Derived Xenograft
PRF: Pulse Repetition Frequency
RCC: Renal Cell Carcinoma
RFA: Radiofrequency Ablation
SCID: Severe Combined Immunodeficiency
SD: Standard Deviation
SEM: Standard Error of the Mean
STS: Soft Tissue Sarcoma
TAM: Tumor Associated Macrophage
xxxv
TDLN: Tumor Draining Lymph Node
tHIFU: Thermal High Intensity Focused Ultrasound
TKX: Telazol, Ketamine, Xylazine
TME: Tumor Microenvironment
US: Ultrasound
VCRO: Veterinary Clinical Trials Network and Clinical Research Office
VMCVM: Virginia-Maryland College of Veterinary Medicine
xxxvi
Attributions for Co-Authored Papers
Alex Simon: Assisted with experimentation, manuscript preparation, and manuscript
review of Chapters 2, 3, 4, 7, and 9.
Allison Zeher: Assisted with experimentation and manuscript review of Chapter 9.
Christopher Bryon, DVM: Assisted with manuscript preparation and manuscript review
for Chapter 4.
David Luyimbazi, MD: Assisted with experiment planning and manuscript review for
Chapters 2, 3, and 7.
Douglas Grider, MD: Provided pathology review for data collection and manuscript
review to Chapters 2, 3, and 7.
Eli Vlaisavljevich, PhD: Assisted with experiment planning, manuscript preparation,
and manuscript review for Chapters 2-9.
Hannah Sheppard: Assisted with experimentation and manuscript review of Chapter 7.
Holly Morrison: Assisted with experimentation and manuscript review of Chapter 6.
Irving C Allen, PhD: Assisted with experiment planning, manuscript preparation, and
manuscript review for Chapters 2-9.
Jaqueline Sereno: Assisted with experimentation and manuscript review of Chapter 9.
Jessica Gannon: Assisted with experimentation, manuscript preparation, and
manuscript review of Chapters 4, 6, 7, and 10. Co-first author of the submitted
manuscripts associated with Chapters 4 and 7.
Joan Vidal-Jove, MD, PhD: Assisted with experiment planning and manuscript review
for Chapters 2 and 7.
xxxvii
Joanne Tuohy, DVM, PhD: Assisted with experiment planning, manuscript preparation,
and manuscript review for Chapters 4.
John H Rossmeisl, DVM: Assisted with manuscript preparation and manuscript review
for Chapter 4.
Kayla Farrell: Assisted with experimentation and manuscript review of Chapter 6.
Kenneth N Aycock: Assisted with experimentation and manuscript review of Chapter 6.
Khan Mohammad Imran: Assisted with experimentation, manuscript preparation, and
manuscript review of Chapters 9.
Kiho Lee, PhD: Assisted with experiment planning, manuscript preparation, and
manuscript review for Chapters 5 and 6.
Kyungjun Uh: Assisted with experimentation, manuscript preparation, and manuscript
review of Chapters 5 and 6.
Lauren Arnold: Assisted with experimentation, manuscript preparation, and manuscript
review of Chapters 4 and 7. Co-first author of the submitted manuscript associated with
Chapter 4.
Margaret Nagai-Singer: Assisted with experimentation, manuscript preparation, and
manuscript review of Chapters 4-7.
Martha M Larson, DVM: Provided radiology assistance for experimentation and
manuscript review of Chapter 7.
Melvin Lorenzo: Assisted with experimentation and manuscript review of Chapter 6.
Michael Edwards, DVM, PhD: Provided radiology assistance for experimentation and
manuscript review of Chapter 7.
xxxviii
Mishal Mendiratta-Lala, MD: Assisted with experiment planning and manuscript review
of Chapter 7.
Natalie Beitel-White: Assisted with experimentation, manuscript preparation, and
manuscript review of Chapters 6 and 9.
Neha Singh: Assisted with experimentation, manuscript preparation, and manuscript
review of Chapters 4 and 7.
Nikolaos Dervisis, DVM: Assisted with experiment planning, manuscript writing, and
manuscript review for Chapter 4.
Peter Weber: Assisted with experimentation and manuscript review of Chapter 2.
Rafael Davalos, PhD: Assisted with experiment planning and manuscript review for
Chapter 6 and 9.
Rebecca Brock, PhD: Assisted with experimentation and manuscript review of
Chapters 6 and 9.
Ruby Hutchison: Assisted with manuscript preparation and review of Chapter 8.
Shawna Klahn, DVM: Assisted with experiment planning, manuscript preparation, and
manuscript review for Chapter 4.
Sherrie Clark-Deener, DVM, PhD: Assisted with experimentation, manuscript
preparation, and manuscript review for Chapters 5 and 6.
Sheryl Coutermarsh-Ott, DVM, PhD: Provided pathology review for data collection
and manuscript review to Chapters 2, 3, 4, 6, 7, and 9.
Sofie Saunier: Assisted with experimentation, manuscript preparation, and manuscript
review of Chapters 2 and 3.
1
Chapter 1. Introduction
2
1.1 Introduction
Cancer is the second leading cause of death world-wide, with multiple tumor types
still facing abysmal 10% or lower 5-year overall survival rates. There are many reasons
attributed to this, but regardless, there is a pressing need to improve treatment options.
The core pillars of cancer therapy are chemotherapy, surgery, radiation,
immunotherapy, and ablation. The most common treatment option for solid tumors
remains surgical resection; however, many patients are not candidates due to tumor
size, location, or disease progression [1-4].
Two cancers that epitomize these problems are pancreatic cancer and
cholangiocarcinoma (CC), a cancer of the bile ducts. Pancreatic cancer is the fourth
leading cause of cancer-related deaths, with a 9% survival rate due to its late diagnosis
and lack of curative treatment options [5, 6]. Although less common, CC also has a low
survival rate of 5-10% [7]. For these malignancies, the standard treatment options are
limited to surgery, chemotherapy, and radiation [7-10]. While surgical resection is the
most curative option for these tumors, many patients do not qualify due to tumor
burden, location, or comorbid disease [2, 11, 12]. Only 20% of patients have pancreatic
tumors that can be surgically removed, and of those patients the cure rate is less than
25% [13]. Only 15% of intrahepatic CC tumors and 56% of distal CC tumors on average
are deemed resectable [14]. For patients with surgically resectable intrahepatic CC, the
5-year survival rate is still only 22-30% [9, 15] This indicates that even with the most
ideal presentation CC still has a low survival rate. In order to address the limitations that
3
are faced in treating these tumor types, there has been recent development of new
ablation therapies and investigation into the potential of immunomodulation to better
fight systemic disease.
1.2 Standard Ablation Procedures for Tumor Debulking and Immunomodulation
Ablation procedures can act as a replacement for or as an adjuvant to surgery. The
most commonly used ablation modalities for treating cancers in abdominal organs are
radiofrequency ablation (RFA), microwave ablation, high-intensity focused ultrasound
(HIFU), cryoablation, and irreversible electroporation (IRE). Of these modalities, RFA,
microwave ablation, and HIFU all utilize thermal mechanisms. RFA is a minimally-
invasive ablation modality that utilizes high-frequency alternating currents to thermally
induce thermal necrosis [16]. Microwave ablation, also minimally-invasive, induces cell
death through electromagnetic microwaves that produce friction and heat [17]. HIFU, a
non-invasive focused ultrasound, ablates cells by depositing ultrasound energy at a
focal point to rapidly increase tissue temperature [18]. While these therapies have
promise, reliance on thermal mechanisms leads to inherent limitations due to the heat
sink effect [19-21] where treatments near vasculature are reduced in efficacy due to the
flow of blood through the treatment region minimizing the efficacy of heat deposition.
This also leads to non-uniform heat dissipation associated with hypervascularization
[22]. An additional limitation to these therapies is the lack of real-time treatment
monitoring, although treatment can be evaluated for success with post-treatment CT
and MR imaging.
4
Alternatively, cryoablation and IRE do not rely on heat dissipation for their
mechanism of ablation. Cryoablation results in tumor cell destruction through ice-crystal
formation as a result of liquid nitrogen or argon gas delivered to the tissue, which can be
monitored in real time with medical imaging [23]. IRE utilizes short, high-voltage
electrical pulses that non-thermally open micro-pores in cell plasma membranes,
inducing cell death, and can be monitored in real-time for changes in tissue electrical
conductivity [24, 25]. Recent advancements have established that non-thermal
therapies have the potential to treat difficult tumors such as pancreatic cancer. For
example, clinical trials have shown that IRE can ablate pancreatic tumors without
damaging nearby critical structures and have had dramatic, positive effects on patient
survival that may be due to the induction of immunomodulatory mechanisms [26-31].
Given cryoablation and IRE’s non-thermal mechanisms, these procedures have been
found to release immunostimulatory molecules and lead to a more immunologically
active tumor microenvironment after treatment [16, 32]. Furthermore, recent studies
comparing the thermal ablation modalities, cryoablation, and IRE have found that the
non-thermal modalities are more potent at stimulating an anti-tumor microenvironment
[33]. However, these procedures still require patients be healthy enough, through co-
morbidities and metastatic disease, for surgery and involve surgical incisions that
increase the possibility of surgery-related injury and infection, including the minimally
invasive percutaneous procedures.
5
1.3 Histotripsy as a Tumor Ablation Therapy
Histotripsy is an emerging non-invasive, non-thermal, image-guided cancer ablation
modality that has recently been approved for its first clinical trial in the United States
(NCT04573881). Histotripsy is a focused ultrasound therapy that utilizes microsecond
through millisecond pulsing regimens to generate controlled cavitation bubble clouds at
the focus, leading to precise ablation of targeted tissues [34] (Fig.1). The rapid
expansion and collapse of cavitation bubbles ablate tissues into acellular debris [35-37].
This process is well
established and is due to the
mechanical nature of the
cavitation-based ablation.
Given the mechanical
mechanism, there is tissue selectivity associated with cavitation ablation. For example,
stronger tissues, with a higher Young’s Modulus, such as vasculature and collecting
ducts are more resistant to damage [38-42]. By utilizing a fully mechanical mechanism,
tissue strength is strongly correlative with dose and susceptibility [43-45]. Studies have
shown that histotripsy is capable of ablating tissues that have a lower concentration of
collagen within the extracellular matrix [39]. This finding, in combination with the
established tissue strength-susceptibility effect, implies increased safety of treating
tumors that frequently have disorganized extracellular matrix components and are
located near stiffer critical structures, such as blood vessels or bile ducts.
Figure 1-1 Histotripsy is a non-invasive ablation method that uses focused ultrasound to generate a cavitation bubble cloud, resulting in the complete destruction of targeted tissues with high precision.
6
Unlike the vast majority of other ablation therapies, histotripsy is non-thermal, and it
is not affected by the heat-sink effect, and ,therefore, remains safe and efficacious for
use near the vasculature [46, 47]. Additional benefits of histotripsy include real-time
imaging feedback with standard imaging (i.e. ultrasound, MRI, CT), highly precise-
millimeter accuracy, and the ability to treat tumors of arbitrary sizes and shapes [40, 48].
An additional benefit of histotripsy is the improved wound healing of ablation zones
compared to other ablation modalities. Histotripsy ablation of healthy rat livers showed
rapid involution of treated volumes, granulation, and growth of healthy hepatocytes
within 28 days with minimal scarring [36]. This healing process is more rapid than what
has been reported for other ablation modalities, such as RFA in humans which has
been reported as “thermal fixation” [49], where the necrotic mass is still present 14
months post-treatment due to thermal denaturation of the tissue leaving it resistant to
being broken down by normal pathways.
In mouse models, histotripsy ablation of hepatic, renal, neuroblastoma, and
melanoma tumors has resulted in significantly increased survival [50-53]. In addition to
debulking tumors, recent evidence has suggested that histotripsy has the capability to
induce a systemic immune response, as evidenced by the attenuation of metastasis and
an improvement in local and distant disease with combination immunotherapy [52, 54,
55]. This effect was shown not only in single animal-tumor treatments, but also showed
abscopal-like decreases in contralateral tumor growth in a separate untreated tumor.
This effect was found to be modest but statistically significant with histotripsy ablation
compared to the insignificant trend in mice treated with irradiation and RFA [52].
Primary studies focused on the use of histotripsy in treating tumors in model organisms
7
have shown that it can be safe and effective; however, there is still plenty of work to be
done before establishing that this therapy will be efficacious in humans [50, 56, 57].
Despite the promise of histotripsy and the many positive results reported in
preclinical studies to date, only a single human clinical trial has been conducted to test
histotripsy for the treatment of cancer [58]. This recent Phase I clinical trial was
conducted with non-curative, palliative intent in patients with multifocal primary and
metastatic liver tumors, with results showing histotripsy could safely target and ablate
targeted liver tumors [58]. To our knowledge, no other human clinical trials of histotripsy
for the treatment of cancer have been conducted.
1.4 Animal Models for Pre-Clinical Testing of Histotripsy
Similar to other focal tumor ablation modalities, one of the primary factors limiting the
rapid translation of histotripsy into the clinic has been the lack of appropriate animal
models for studying the safety, effectiveness, and optimization for the treatment of a
variety of cancer types. Previous efforts to develop histotripsy for specific oncology
applications have been split into separate studies testing human-scale and miniaturized
rodent-scale systems.
Studies testing human-scale histotripsy systems for ablating tissues is normally
conducted in healthy, large animal models that have similar anatomy and physiology to
human patients, primarily pigs [48, 59-61]. For example, the same tumor ablation
systems can typically be utilized, similar clinical/surgical techniques can be refined, and
many large animal models accurately recapitulate human anatomical and physiological
8
features. This is critical for imaging and ultrasound-based modalities such as histotripsy.
Although early pre-clinical studies utilizing pigs has focused on healthy animals, the
development of novel cancer porcine models is expanding the potential for pre-clinical
work. With the rise of technologies capable of inducing genetic modifications in large
mammals, such as CRISPRs and TALONs, there is an interest in the pig as a
spontaneous cancer model [62-64]. As an alternative strategy, research teams have
focused on immunocompromised pig models, where deletions in the RAG2 and IL2RG
genes have generated animals with similar characteristics to NOD scid gamma (NSG)
mice, which are susceptible to human tissue engraftment [65]. Similar to NSG mice,
human tumor cell lines and patient-derived xenograft (PDX) tissue can be engrafted [66,
67]. This allows a highly predictable tumor that can be surgically placed subcutaneously
or orthotopically in a human-sized animal with similar anatomy and physiology. In
general, pigs have yet to play a significant role in experimental oncology [62].
These large animal debulking studies are typically complemented by separately
testing histotripsy tumor ablation in rodents (mice, rats, and rabbits) using customized
small animal devices that allow histotripsy to be applied in this setting. Albeit, these
studies do not use the systems or acoustic parameters that will ultimately be developed
for use in humans in order to more rapidly investigate the biological and immunological
effects of ablation [52, 54, 56, 68]. Murine models are the dominant in vivo model in the
biomedical engineering and cancer biology fields and are essential in acquiring
fundamental data. In typical mouse models used to study tumor ablation modalities, the
vast majority of studies are mouse or human cell line based. While these models offer
excellent reproducibility and standardization, they rarely recapitulate the clinical, in situ,
9
microenvironment found in patients or critical biological hallmarks of the tumor of origin.
Likewise, the lack of translational acoustic windows, limited imaging, lack of precision,
and lack of physiological relevance are all major limitations of small animal models in
the development of histotripsy and other focused ultrasound modalities.
Beyond induced tumor models, veterinary cancer patients offer unique
translational research opportunities for cancer imaging and medical device
development. Similar to humans, companion animals are genetically diverse and are
exposed to similar environments. Comparative oncology studies have revealed
significant similarities between many canine and human cancers, and over the past two
decades canines have been used for an increasing number of pre-clinical pharmacology
studies [69-77]. The spontaneous nature of canine cancers, their biological similarities
to human cancers, and the anatomic similarities between dogs and people offer highly
relevant clinical conditions for device development.
1.5 Outline of this Dissertation
This dissertation consists of three parts. Part One of this dissertation (Chapters
2-3) focuses on determining the safety and dosages for using histotripsy to treat
stronger hepatopancreatobiliary tumors using excised tissues and patient derived
xenograft (PDX) mice. Chapter 2 focuses on dosimetry studies of CC tumors, to
determine the feasibility of using histotripsy to treat dense, highly fibrotic tumors. For
Chapter 4, PDX mice bearing CC tumors were treated with histotripsy and survived to
determine the feasibility of a full ablation without debilitating off target damage.
10
Part Two (Chapters 4-7) moves onto more complex models, using large animals
and veterinary patients to establish application protocols for additional tumor types.
Chapter 5 demonstrates the pros and cons of utilizing histotripsy to treat a wide range of
tumors in murine and porcine models as well as veterinary canine patients. Chapter 6
introduces an immunocompromised porcine model that can support the growth of
human tumors. Chapter 7 utilizes the newly established xenograft porcine model to
grow human pancreatic tumors and compares the physiology of these tumors and pigs
to humans, with a pilot study showing the efficacy of using these tumors for IRE
ablation. In Chapter 8, healthy porcine pancreases are ablated in vivo to determine the
safety and feasibility of using histotripsy to treat targets within the pancreas.
Part Three (Chapters 8-9) shifts to utilizing immunocompetent mouse models to
investigate the immunological effects of histotripsy in pancreatic adenocarcinoma. In
Chapter 9, the current body of literature on histotripsy’s role in modulating the immune
system locally and systemically is reviewed. Chapter 10 uses the Pan02 pancreatic
adenocarcinoma subcutaneous model in C57/Bl6 mice to investigate the release of
damage associate molecular patterns and potential antigens by histotripsy compared to
established ablation modalities as well as the shift in the local tumor microenvironment.
11
1.6 References
[1] A. S. Befeler and A. M. Di Bisceglie, "Hepatocellular carcinoma: diagnosis and
treatment," Gastroenterology, vol. 122, no. 6, pp. 1609-1619, 2002.
[2] E. G. Chiorean and A. L. Coveler, "Pancreatic cancer: optimizing treatment
options, new, and emerging targeted therapies," Drug design, development and
therapy, vol. 9, pp. 3529-45, 2015, doi: 10.2147/DDDT.S60328.
[3] C. Armengol, M. R. Sarrias, and M. Sala, "Hepatocellular carcinoma: present and
future," Medicina Clínica (English Edition), vol. 150, no. 10, pp. 390-397, 2018.
[4] J. J. Hsieh et al., "Renal cell carcinoma," Nature reviews Disease primers, vol. 3,
p. 17009, 2017.
[5] R. L. Siegel, K. D. Miller, and A. Jemal, "Cancer statistics, 2017," CA: a cancer
journal for clinicians, vol. 67, no. 1, pp. 7-30, 2017.
[6] R. L. Siegel, K. D. Miller, and A. Jemal, "Cancer statistics, 2020," CA: a cancer
journal for clinicians, vol. 70, no. 1, pp. 7-30, 2020.
[7] E. U. Cidon, "Resectable cholangiocarcinoma: reviewing the role of adjuvant
strategies," Clinical Medicine Insights: Oncology, vol. 10, p. CMO. S32821, 2016.
[8] H. Manuel, "Pancreatic cancer," New England Journal of Medicine, vol. 362, no.
17, 2010.
[9] M. N. Mavros, K. P. Economopoulos, V. G. Alexiou, and T. M. Pawlik, "Treatment
and prognosis for patients with intrahepatic cholangiocarcinoma: systematic
review and meta-analysis," JAMA surgery, vol. 149, no. 6, pp. 565-574, 2014.
12
[10] J. Liu et al., "Prognostic Factors and Treatment Strategies for Intrahepatic
Cholangiocarcinoma from 2004 to 2013: Population-Based SEER Analysis,"
Transl Oncol, vol. 12, no. 11, pp. 1496-1503, Nov 2019, doi:
10.1016/j.tranon.2019.05.020.
[11] M. C. Jansen, R. van Hillegersberg, R. A. Chamuleau, O. M. van Delden, D. J.
Gouma, and T. M. van Gulik, "Outcome of regional and local ablative therapies
for hepatocellular carcinoma: a collective review," Eur J Surg Oncol, vol. 31, no.
4, pp. 331-47, May 2005, doi: 10.1016/j.ejso.2004.10.011.
[12] L. Rossi et al., "Current approach in the treatment of hepatocellular carcinoma,"
World J Gastrointest Oncol, vol. 2, no. 9, pp. 348-59, Sep 15 2010, doi:
10.4251/wjgo.v2.i9.348.
[13] S. Gillen, T. Schuster, C. M. Zum Büschenfelde, H. Friess, and J. Kleeff,
"Preoperative/neoadjuvant therapy in pancreatic cancer: a systematic review and
meta-analysis of response and resection percentages," PLoS medicine, vol. 7,
no. 4, 2010.
[14] D. M. Nagorney, J. H. Donohue, M. B. Farnell, C. D. Schleck, and D. M. Ilstrup,
"Outcomes after curative resections of cholangiocarcinoma," Archives of surgery,
vol. 128, no. 8, pp. 871-878, 1993.
[15] G. Spolverato et al., "Can hepatic resection provide a long‐term cure for patients
with intrahepatic cholangiocarcinoma?," Cancer, vol. 121, no. 22, pp. 3998-4006,
2015.
13
[16] O. Kepp, A. Marabelle, L. Zitvogel, and G. Kroemer, "Oncolysis without viruses—
inducing systemic anticancer immune responses with local therapies," Nature
Reviews Clinical Oncology, pp. 1-16, 2019.
[17] C. J. Simon, D. E. Dupuy, and W. W. Mayo-Smith, "Microwave ablation:
principles and applications," Radiographics, vol. 25, no. suppl_1, pp. S69-S83,
2005.
[18] Y. Keisari, "Tumor abolition and antitumor immunostimulation by physico-
chemical tumor ablation," Front Biosci (Landmark Ed), vol. 22, pp. 310-47, 2017.
[19] S. A. Curley, "Radiofrequency ablation of malignant liver tumors," The oncologist,
vol. 6, no. 1, pp. 14-23, 2001.
[20] D. S. Lu et al., "Influence of large peritumoral vessels on outcome of
radiofrequency ablation of liver tumors," Journal of vascular and interventional
radiology, vol. 14, no. 10, pp. 1267-1274, 2003.
[21] J. A. Marrero and S. Pelletier, "Hepatocellular carcinoma," Clin Liver Dis, vol. 10,
no. 2, pp. 339-51, ix, May 2006, doi: 10.1016/j.cld.2006.05.012.
[22] T. Livraghi, L. Solbiati, M. F. Meloni, G. S. Gazelle, E. F. Halpern, and S. N.
Goldberg, "Treatment of focal liver tumors with percutaneous radio-frequency
ablation: complications encountered in a multicenter study," Radiology, vol. 226,
no. 2, pp. 441-451, 2003.
[23] J. P. Erinjeri and T. W. Clark, "Cryoablation: mechanism of action and devices,"
Journal of Vascular and Interventional Radiology, vol. 21, no. 8, pp. S187-S191,
2010.
14
[24] R. V. Davalos, L. Mir, and B. Rubinsky, "Tissue ablation with irreversible
electroporation," Annals of biomedical engineering, vol. 33, no. 2, p. 223, 2005.
[25] R. V. Davalos, B. Rubinsky, and D. M. Otten, "A feasibility study for electrical
impedance tomography as a means to monitor tissue electroporation for
molecular medicine," IEEE Transactions on Biomedical Engineering, vol. 49, no.
4, pp. 400-403, 2002.
[26] R. C. Martin et al., "Treatment of 200 locally advanced (stage III) pancreatic
adenocarcinoma patients with irreversible electroporation: safety and efficacy,"
Annals of surgery, vol. 262, no. 3, pp. 486-494, 2015.
[27] H. J. Scheffer et al., "Ablation of locally advanced pancreatic cancer with
percutaneous irreversible electroporation: results of the phase I/II PANFIRE
study," Radiology, vol. 282, no. 2, pp. 585-597, 2017.
[28] M. P. Belfiore et al., "Percutaneous CT-guided irreversible electroporation
followed by chemotherapy as a novel neoadjuvant protocol in locally advanced
pancreatic cancer: our preliminary experience," International Journal of Surgery,
vol. 21, pp. S34-S39, 2015.
[29] H. J. Scheffer et al., "Irreversible electroporation of locally advanced pancreatic
cancer transiently alleviates immune suppression and creates a window for
antitumor T cell activation," Oncoimmunology, vol. 8, no. 11, p. 1652532, 2019.
[30] R. C. Martin et al., "Irreversible electroporation in locally advanced pancreatic
cancer: a call for standardization of energy delivery," Journal of Surgical
Oncology, vol. 114, no. 7, pp. 865-871, 2016.
15
[31] N. Beitel-White, R. C. G. Martin, Y. Li, R. M. Brock, I. C. Allen, and R. V.
Davalos, "Real-time prediction of patient immune cell modulation during
irreversible electroporation therapy," (in eng), Sci Rep, vol. 9, no. 1, p. 17739,
Nov 28 2019, doi: 10.1038/s41598-019-53974-w.
[32] R. M. Brock, N. Beitel-White, R. V. Davalos, and I. C. Allen, "Starting a Fire
Without Flame: The Induction of Cell Death and Inflammation in Electroporation-
Based Tumor Ablation Strategies," Frontiers in Oncology, vol. 10, 2020.
[33] Q. Shao et al., "Engineering T cell response to cancer antigens by choice of focal
therapeutic conditions," International Journal of Hyperthermia, vol. 36, no. 1, pp.
130-138, 2019.
[34] K. B. Bader, E. Vlaisavljevich, and A. D. Maxwell, "For whom the bubble grows:
physical principles of bubble nucleation and dynamics in histotripsy ultrasound
therapy," Ultrasound in medicine & biology, 2019.
[35] A. R. Smolock et al., "Robotically Assisted Sonic Therapy as a Noninvasive
Nonthermal Ablation Modality: Proof of Concept in a Porcine Liver Model,"
Radiology, vol. 287, no. 2, pp. 485-493, May 2018, doi:
10.1148/radiol.2018171544.
[36] E. Vlaisavljevich et al., "Non-invasive ultrasound liver ablation using histotripsy:
Chronic study in an in vivo rodent model," Ultrasound in medicine & biology, vol.
42, no. 8, pp. 1890-1902, 2016.
[37] E. Vlaisavljevich et al., "Non-invasive liver ablation using histotripsy: preclinical
safety study in an in vivo porcine model," Ultrasound in medicine & biology, vol.
43, no. 6, pp. 1237-1251, 2017.
16
[38] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," Physics in Medicine & Biology, vol. 59, no. 2, p. 253, 2013.
[39] E. Vlaisavljevich, Z. Xu, A. Arvidson, L. Jin, W. Roberts, and C. Cain, "Effects of
thermal preconditioning on tissue susceptibility to histotripsy," Ultrasound in
medicine & biology, vol. 41, no. 11, pp. 2938-2954, 2015.
[40] T. L. Hall, C. R. Hempel, K. Wojno, Z. Xu, C. A. Cain, and W. W. Roberts,
"Histotripsy of the prostate: dose effects in a chronic canine model," Urology, vol.
74, no. 4, pp. 932-937, 2009.
[41] A. Lake, Z. Xu, J. Wilkinson, C. Cain, and W. Roberts, "Renal ablation by
histotripsy—does it spare the collecting system?," The Journal of urology, vol.
179, no. 3, pp. 1150-1154, 2008.
[42] C. R. Hempel, T. L. Hall, C. A. Cain, J. B. Fowlkes, Z. Xu, and W. W. Roberts,
"Histotripsy fractionation of prostate tissue: local effects and systemic response
in a canine model," The Journal of urology, vol. 185, no. 4, pp. 1484-1489, 2011.
[43] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," (in eng), Phys Med Biol, vol. 59, no. 2, pp. 253-70, Jan 20 2014, doi:
10.1088/0031-9155/59/2/253.
[44] E. Vlaisavljevich, A. Maxwell, M. Warnez, E. Johnsen, C. Cain, and Z. Xu,
"Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
mechanical properties," (in eng), IEEE Transactions on Ultrasonics,
17
Ferroelectrics, and Frequency Control, vol. 61, no. 2, pp. 341-52, Feb 2014, doi:
10.1109/TUFFC.2014.6722618.
[45] V. A. Khokhlova et al., "Histotripsy methods in mechanical disintegration of
tissue: Towards clinical applications," International journal of hyperthermia, vol.
31, no. 2, pp. 145-162, 2015.
[46] L. Thanos, S. Mylona, P. Galani, M. Pomoni, A. Pomoni, and I. Koskinas,
"Overcoming the heat-sink phenomenon: successful radiofrequency thermal
ablation of liver tumors in contact with blood vessels," Diagnostic and
Interventional Radiology, vol. 14, no. 1, p. 51, 2008.
[47] K. Pillai et al., "Heat sink effect on tumor ablation characteristics as observed in
monopolar radiofrequency, bipolar radiofrequency, and microwave, using ex vivo
calf liver model," Medicine, vol. 94, no. 9, 2015.
[48] C. M. Smolock AR, Vlaisavljevich E, Gendron-Fitzpatrick A, Green C, Ziemlewicz
TJ and Lee FT, "Robotically Assisted Sonic Therapy as a Non-Invasive Non-
Thermal Ablation Modality: Proof of Concept in a Porcine Liver Model,"
Radiology, vol. 287, no. 2, pp. 485-93, 2018.
[49] J. E. Coad, K. Kosari, A. Humar, and T. D. Sielaff, "Radiofrequency ablation
causes ‘thermal fixation’of hepatocellular carcinoma: a post‐liver transplant
histopathologic study," Clinical transplantation, vol. 17, no. 4, pp. 377-384, 2003.
[50] G. R. Schade, "Evaluation of the Systemic Response to Boiling Histotripsy
Treatment for Renal Carcinoma," 2018.
18
[51] T. Worlikar et al., "Non-Invasive Orthotopic Liver Tumor Ablation Using
Histotripsy in an in Vivo Rodent Hepatocellular Carcinoma (HCC) Model,"
presented at the Society of Interventional Oncology, Boston, MA, USA, 2019.
[52] S. Qu et al., "Non-thermal histotripsy tumor ablation promotes abscopal immune
responses that enhance cancer immunotherapy," Journal for ImmunoTherapy of
Cancer, vol. 8, no. 1, 2020.
[53] A. Eranki et al., "High-Intensity Focused Ultrasound (HIFU) Triggers Immune
Sensitization of Refractory Murine Neuroblastoma to Checkpoint Inhibitor
Therapy," Clinical Cancer Research, vol. 26, no. 5, pp. 1152-1161, 2020.
[54] T. Worlikar et al., "Histotripsy for Non-Invasive Ablation of Hepatocellular
Carcinoma (HCC) Tumor in a Subcutaneous Xenograft Murine Model," Conf Proc
IEEE Eng Med Biol Soc, vol. 2018, pp. 6064-6067, Jul 2018, doi:
10.1109/EMBC.2018.8513650.
[55] G. R. Schade, Y.-N. Wang, S. D'Andrea, J. H. Hwang, W. C. Liles, and T. D.
Khokhlova, "Boiling Histotripsy Ablation of Renal Cell Carcinoma in the Eker Rat
Promotes a Systemic Inflammatory Response," Ultrasound in medicine &
biology, vol. 45, no. 1, pp. 137-147, 2019.
[56] T. Worlikar et al., "Non-invasive liver cancer ablation using histotripsy in an in
vivo murine hepatocellular carcinoma (HCC) model," in Ultrasonics Symposium
(IUS), 2017 IEEE International, 2017: IEEE, pp. 1-1.
[57] K. J. Pahk et al., "Boiling Histotripsy-induced Partial Mechanical Ablation
Modulates Tumour Microenvironment by Promoting Immunogenic Cell Death of
Cancers," Scientific reports, vol. 9, no. 1, pp. 1-12, 2019.
19
[58] S. X. Vidal Jove J, Vlaisavljevich E, Cannata J, Duryea A, Miller R, Lee F and
Ziemlewicz T, "Phase I Study of Safety and Efficacy of Hepatic Histotripsy:
Preliminary Results of First in Man Experience with Robotically-Assisted Sonic
Therapy," in International Society of Therapeutic Ultrasound, Barcelona, Spain,
2019.
[59] V. E. Kim Y, Owens GE, Allen SP, Cain CA and Xu Z, "In vivo transcostal
histotripsy therapy without aberration correction," Phys Med Biol, vol. 59, pp.
2553-68, 2014.
[60] E. Vlaisavljevich et al., "Image-guided non-invasive ultrasound liver ablation
using histotripsy: feasibility study in an in vivo porcine model," Ultrasound Med
Biol, vol. 39, no. 8, pp. 1398-409, Aug 2013, doi:
10.1016/j.ultrasmedbio.2013.02.005.
[61] E. Vlaisavljevich et al., "Non-Invasive Liver Ablation Using Histotripsy: Preclinical
Safety Study in an In Vivo Porcine Model," Ultrasound Med Biol, vol. 43, no. 6,
pp. 1237-1251, Jun 2017, doi: 10.1016/j.ultrasmedbio.2017.01.016.
[62] T. Flisikowska, A. Kind, and A. Schnieke, "Pigs as models of human cancers,"
Theriogenology, vol. 86, no. 1, pp. 433-7, Jul 1 2016, doi:
10.1016/j.theriogenology.2016.04.058.
[63] K. M. Schachtschneider et al., "The Oncopig Cancer Model: An Innovative Large
Animal Translational Oncology Platform," Front Oncol, vol. 7, p. 190, 2017, doi:
10.3389/fonc.2017.00190.
[64] L. B. Schook et al., "A Genetic Porcine Model of Cancer," PLoS One, vol. 10, no.
7, p. e0128864, 2015, doi: 10.1371/journal.pone.0128864.
20
[65] K. Lee, K. Farrell, and K. Uh, "Application of genome-editing systems to enhance
available pig resources for agriculture and biomedicine," (in eng), Reprod Fertil
Dev, vol. 32, no. 2, pp. 40-49, Jan 2019, doi: 10.1071/rd19273.
[66] J. J. Tentler et al., "Patient-derived tumour xenografts as models for oncology
drug development," (in eng), Nature reviews. Clinical oncology, vol. 9, no. 6, pp.
338-50, Apr 17 2012, doi: 10.1038/nrclinonc.2012.61.
[67] R. M. Brock et al., "Patient derived xenografts expand human primary pancreatic
tumor tissue availability for ex vivo irreversible electroporation testing," Frontiers
in Oncology, vol. 10, 2020.
[68] E. Vlaisavljevich et al., "Non-Invasive Ultrasound Liver Ablation Using Histotripsy:
Chronic Study in an In Vivo Rodent Model," Ultrasound Med Biol, vol. 42, no. 8,
pp. 1890-902, Aug 2016, doi: 10.1016/j.ultrasmedbio.2016.03.018.
[69] S. S. Pinho, S. Carvalho, J. Cabral, C. A. Reis, and F. Gartner, "Canine tumors: a
spontaneous animal model of human carcinogenesis," Transl Res, vol. 159, no.
3, pp. 165-72, Mar 2012, doi: 10.1016/j.trsl.2011.11.005.
[70] J. D. Schiffman and M. Breen, "Comparative oncology: what dogs and other
species can teach us about humans with cancer," Philos Trans R Soc Lond B
Biol Sci, vol. 370, no. 1673, Jul 19 2015, doi: 10.1098/rstb.2014.0231.
[71] M. Paoloni et al., "Defining the Pharmacodynamic Profile and Therapeutic Index
of NHS-IL12 Immunocytokine in Dogs with Malignant Melanoma," PLoS One, vol.
10, no. 6, p. e0129954, 2015, doi: 10.1371/journal.pone.0129954.
[72] M. Paoloni et al., "Prospective molecular profiling of canine cancers provides a
clinically relevant comparative model for evaluating personalized medicine
21
(PMed) trials," PLoS One, vol. 9, no. 3, p. e90028, 2014, doi:
10.1371/journal.pone.0090028.
[73] M. Paoloni, S. Lana, D. Thamm, C. Mazcko, and S. Withrow, "The creation of the
Comparative Oncology Trials Consortium Pharmacodynamic Core: Infrastructure
for a virtual laboratory," Vet J, vol. 185, no. 1, pp. 88-9, Jul 2010, doi:
10.1016/j.tvjl.2010.04.019.
[74] M. C. Paoloni et al., "Rapamycin pharmacokinetic and pharmacodynamic
relationships in osteosarcoma: a comparative oncology study in dogs," PLoS
One, vol. 5, no. 6, p. e11013, Jun 08 2010, doi: 10.1371/journal.pone.0011013.
[75] I. Gordon, M. Paoloni, C. Mazcko, and C. Khanna, "The Comparative Oncology
Trials Consortium: using spontaneously occurring cancers in dogs to inform the
cancer drug development pathway," PLoS Med, vol. 6, no. 10, p. e1000161, Oct
2009, doi: 10.1371/journal.pmed.1000161.
[76] M. C. Paoloni et al., "Launching a novel preclinical infrastructure: comparative
oncology trials consortium directed therapeutic targeting of TNFalpha to cancer
vasculature," PLoS One, vol. 4, no. 3, p. e4972, 2009, doi:
10.1371/journal.pone.0004972.
[77] C. A. London et al., "Preclinical evaluation of the novel, orally bioavailable
Selective Inhibitor of Nuclear Export (SINE) KPT-335 in spontaneous canine
cancer: results of a phase I study," PLoS One, vol. 9, no. 2, p. e87585, 2014, doi:
10.1371/journal.pone.0087585.
22
Chapter 2. Histotripsy for the Treatment of Cholangiocarcinoma Liver Tumors: In Vivo Feasibility and Ex Vivo Dosimetry Study
Alissa Hendricks, Peter Weber, Alex Simon, Sofie Saunier, Sheryl Coutermarsh-Ott, Douglas Grider, Joan Vidal Jove, Irving Coy Allen, David Luyimbazi, Eli Vlaisavljevich
This chapter is excerpted from a manuscript published in IEEE Transactions in UFFC.
2021 IEEE. Reprinted, with permission, from: A. Hendricks-Wenger et al., "Histotripsy for the Treatment of Cholangiocarcinoma Liver Tumors: In Vivo Feasibility and Ex Vivo Dosimetry Study," in IEEE Transactions on Ultrasonics, Ferroelectrics, and Frequency
Control, doi: 10.1109/TUFFC.2021.3073563.
23
2.1 Abstract
Histotripsy is a non-invasive, non-ionizing, and non-thermal focused ultrasound
ablation method that is currently being developed for the treatment of liver cancer.
Promisingly, histotripsy has been shown for ablating primary (hepatocellular carcinoma,
HCC) and metastatic (colorectal liver metastasis, CLM) liver tumors in preclinical and
early clinical studies. The feasibility of treating cholangiocarcinoma (CC), a less
common primary liver tumor that arises from the bile ducts, has not been explored
previously. Given that prior work has established that histotripsy susceptibility is based
on tissue mechanical properties, there is a need to explore histotripsy as a treatment for
CC due to their dense fibrotic stromal components. In this work, we first investigated the
feasibility of histotripsy for ablating CC tumors in vivo in a patient-derived xenograft
mouse model. The results showed that histotripsy could generate CC tumor ablation
using a 1 MHz small animal histotripsy system with treatment doses of 250, 500, and
1000 pulses/point. A second set of experiments compared the histotripsy doses
required to ablate CC tumors to HCC and CLM tumors ex vivo. For this, human tumor
samples were harvested after surgery and treated ex vivo with a 700 kHz clinical
histotripsy transducer. Results demonstrated significantly higher treatment doses were
required to ablate CC and CLM tumors compared to HCC, with the highest treatment
dose required for CC tumors. Overall, the results of this study suggest that histotripsy
has the potential to be used for the ablation of CC tumors while also highlighting the
need for tumor-specific treatment strategies.
24
2.2 Keywords:
Focused Ultrasound, Histotripsy, Liver Cancer, Cholangiocarcinoma, Hepatocellular
Carcinoma, Liver Metastasis
25
2.3 Introduction
More than 800,000 people are diagnosed with a malignancy originating from or
metastatic to the liver every year. Of these, 42,810 cases are projected for the U.S. in
2020 [1]. These malignant liver tumors include hepatocellular carcinoma (HCC), liver
metastasis (LM), and cholangiocarcinoma (CC). Although surgical resection remains the
mainstay of liver tumor intervention, less than 25% of patients are candidates for
resection due to tumor burden, location, or comorbid disease [2, 3]. For example, a prior
study found that only 15% of intrahepatic CC tumors and 56% of distal tumors were
deemed resectable [4].
Minimally-invasive ablation methods have shown some success in treating liver
tumors but have limitations due to the thermal mechanism, including lack of precision
and high complication rates when treating near critical structures [5-14]. Histotripsy is a
non-invasive, non-thermal, and image-guided focused ultrasound ablation method that
ablates tissue through the generation of acoustic cavitation at the transducer focus [15-
19]. Histotripsy uses microsecond long, high-pressure pulses to generate a histotripsy
bubble cloud inside of the targeted tissue [20-24]. The rapid expansion and collapse of
the cavitation bubbles results in complete disruption of the tissue [25]. As a non-thermal
ablation modality, histotripsy has shown the potential to overcome the limitations
associated with thermal ablation. For instance, hepatic histotripsy has been shown to
produce consistent and complete ablation in highly vascular regions of the liver [18, 26].
Histotripsy has also shown the ability to preserve vital structures such as major vessels,
26
bile ducts, and nerves within the ablation zone [18, 27]. Preclinical in vivo studies of
hepatic histotripsy in healthy porcine and rodent models have demonstrated that
histotripsy can generate clinically relevant ablation zones with sharp boundaries (<1mm)
between treated and healthy liver tissue [15, 17, 18, 26, 27]. Additional studies have
shown the in vivo feasibility of treating HCC tumors in rodent models using histotripsy
[28, 29]. Based on these successful preclinical studies, a Phase I clinical trial in patients
with liver cancer was recently conducted to demonstrate the feasibility of hepatic
histotripsy [16]. Together, these studies have established the feasibility and overall
safety profile of hepatic histotripsy.
Although one patient with CC was reportedly treated as part of the clinical trial
mentioned above, there have been limited studies investigating the potential of
histotripsy for treating CC tumors. CC tumors pose a unique challenge for ablation
modalities due to their common location near vital critical structures [30]. The vessel
and duct sparing features of hepatic histotripsy are promising for the treatment of CC
located near these structures. However, given prior studies showing fibrous tissues with
higher stiffness and mechanical strength are more resistant to histotripsy, it is also
expected that CC tumors may pose a unique challenge for histotripsy. This hypothesis
is based on the CC tumor’s dense desmoplastic fibrotic composition and higher stiffness
that has been reported previously [31-33]. For instance, prior work has found the
average Young’s modulus of CC tumors to be 56.9±25.6 kPa, which was significantly
higher than HCC (14.86±10.0 kPa) and LM (28.8±16.0 kPa) tumors [34]. Given these
mechanical profiles, we hypothesize that histotripsy will require higher treatment doses
to ablate CC tumors compared to LM and HCC.
27
In this study, we investigate the feasibility of using histotripsy for the treatment of
CC tumors. To test histotripsy for CC ablation, this study was split into two parts. In part
one, we investigate the in vivo feasibility of histotripsy CC ablation in a subcutaneous
patient-derived xenograft (PDX) CC tumor model. In part two, we perform a comparative
dosimetry study for treating ex vivo human HCC, LM, and CC tumors with histotripsy to
determine the differences in treatment dose requirements for ablating each tumor type
with histotripsy.
2.4 Procedures
2.4.1 Histotripsy Systems
Two separate histotripsy systems were used in this study for the small animal and
large animal in vivo histotripsy studies. Small animal in vivo experiments were
conducted using a custom 8-element 1MHz histotripsy transducer with a geometric focal
distance of 36 mm, an aperture size of 52.7 mm, and an f-number of 0.68 (Fig.1B). The
transducer was driven via a custom high-voltage pulser designed to generate short
therapy pulses of <2 cycles controlled by a field-programmable gate array (FPGA)
board (Altera DE0-Nano Terasic Technology, Dover, DE, USA) programmed for
histotripsy therapy pulsing. The transducer was positioned in a tank of degassed water
beneath a custom-designed mouse surgical stage (Fig.1) and attached to a computer-
guided 3-D positioning system with a 0.05 mm motor resolution to control the automated
volumetric treatments. A linear ultrasound imaging probe with a frequency range of 10-
28
18 MHz (L18-10L30H-4, Telemed, Lithuania, EU) was coaxially aligned inside the
transducer for treatment guidance and monitoring [28, 29]. The transducer was powered
by a high voltage DC power supply (GENH750W, TDK-Lambda), and the system was
controlled using a custom user-interface operated through MATLAB (MathWorks).
For the ex vivo experiments, histotripsy was applied using a 36-element 700 kHz
clinical histotripsy transducer and associated driving systems (HistoSonics, Inc) that
was previously used in a human clinical trial for the treatment of benign prostatic
hyperplasia [35] and multiple previous pre-clinical studies [36, 37]. This system is the
same frequency as the clinical histotripsy system used in the recent Phase I human liver
cancer trial [16] and was thus chosen for this study. The transducer that consists of a
concave circular aperture with a small inlet on the perimeter to allow for histotripsy
treatments to be guided by a trans-rectal imaging probe (Note: this imaging probe was
not used in the current study [36, 37]. The transducer has a geometric focal distance of
11 cm and FWHM dimensions at a geometric focus of 4.0 mm, 4.0 mm, and 15.0 mm in
the transverse, elevational, and axial dimensions, respectively. The transducer was
powered by a high voltage DC power supply (GENH750W, TDK-Lambda), and the
system was controlled using a custom user-interface called the Histotripsy Service Tool
(HistoSonics, Inc.) to produce 5-cycle histotripsy pulses [24, 38].
2.4.2 Pressure Calibration
Focal pressure waveforms for the 1 MHz and 700 kHz transducers were
measured by a high sensitivity rod hydrophone (HNR-0500, Onda Corporation,
29
Sunnyvale, CA, USA) and a custom-built fiber optic probe hydrophone (FOPH) [39, 40]
in degassed water at the focal point of each transducer. Focal pressures were directly
measured with the FOPH up to a peak negative pressure (p-) of ~16MPa for the 700kHz
transducer and ~20 MPa for the 1MHz transducer. Pressures were not directly
measured beyond these limits due to cavitation on the tip of the hydrophone probe.
Waveforms at higher p- were estimated using linear summation of the pressure
waveforms directly measured from four sub-apertures of the array for the 700 kHz
transducer, and two sub-apertures for the 1MHz transducer. This method showed close
agreement between directly measured pressures and the cavitation threshold seen
experimentally [41].All waveforms were measured using a Tektronix TBS2000 series
oscilloscope at a sample rate of 500MS/s, with the waveforms averaged over 128
pulses and recorded in MATLAB.
2.4.3 Histotripsy In Vivo CC Ablation Procedure
The feasibility of histotripsy for CC tumor ablation was tested in vivo using a
patient-derived xenograft (PDX) CC mouse model. Twelve female NSG mice with
subcutaneous flank PDX CC tumors were ordered from and prepared by Jackson
Laboratory (Bar Harbor, Maine, USA). Through a process previously described for
pancreatic PDX tumors [42], a human CC tumor was collected from a patient and
propagated as a heterogeneous tumor in immunocompromised NSG mice. Once
received, animals were monitored three times weekly under Virginia Tech IACUC
protocols. At each check, general health was monitored and tumor diameter was
30
determined by taking two perpendicular measurements, one of which was the widest
point of the tumor. A single tumor diameter was calculated as the square root of the
product of the two measurements. The mice underwent histotripsy treatment when their
tumors reach the targeted treatment size of ~5mm in diameter. Before treatment, fur
was removed over the tumor and the surrounding area by applying Nair (Naircare,
Ewing, NJ, USA) for 1-2 minutes and removed by wiping off with a wet paper towel. For
treatment, mice were anesthetized with isoflurane (1.5 L/min oxygen flow rate with 1.0-
2.5% isoflurane) and placed prone on the subject stage (Fig.1) with their tumor
submerged underwater. Animals’ respiratory rate and rhythm were monitored during
Figure 2-1 In Vivo Ablation Setup. (A) Histotripsy murine experiments were conducted using a custom 1MHz small animal histotripsy system. (B) Subject’s tumors were submerged in degassed water at the transducer focus in order for (C) histotripsy to be applied to the tumor non-invasively guided by real-time ultrasound imaging.
31
treatment by trained personnel.
The overall setup for the in vivo treatment is illustrated in Figure 1. This custom
system, which has been used in previous small animal in vivo histotripsy studies [17,
28, 29], consists of the 1 MHz histotripsy transducer with co-axially aligned ultrasound
imaging positioned vertically underneath a small animal surgical stage inside a heated
water tank that allows for acoustic coupling. A second water tank is connected to this
primary tank to circulate warmed, degassed water at a constant temperature throughout
the procedures. Before treatment, the water tanks were filled with water and degassed
for approximately 1 hour. A water heater was used to maintain the water temperature at
37+4oC throughout treatment. A small animal platform containing a hole in the center
was located on top of the tank for an anesthetized mouse to be placed with the mouse’s
flank containing the tumor submerged in the water at the transducer focus. The
transducer and a co-axially aligned imaging probe were connected to a 3D motorized
positioning system that was used to apply an automated volumetric ablation to a
targeted volume within the tumor. This custom volumetric ablation system was designed
to allow the user to outline the desired tumor and treatment regions before starting
treatment. The treatment volume was then divided into a 3D grid of points, and
histotripsy was raster-scanned through a series of treatment points spaced by 1.5 mm
in the axial direction and 0.5 mm in the lateral directions. While all the tumors were
treated at different sizes due to delayed growth and planning in between treatments, the
goal was to treat tumors when they averaged 5 mm in diameter. For a 5 mm diameter
tumor, the spacing resulted in a treatment with 1382 treatment points, a dwell time of 2
seconds, and a 46-minute treatment. This treatment spacing was determined based on
32
preliminary experiments to ensure overlap between adjacent bubble clouds. Three
separate treatment doses of 250, 500, and 1000 pulses/point were tested to determine
the feasibility of histotripsy for treating CC tumors in vivo, with the results compared to
an untreated group (n=3/group). Treatment dose was determined based on prior small
animal studies that utilized 50-500 pulses to treat healthy liver and HCC tumors in
addition to the hypothesis that CC tumors would require a high dose given its higher
mechanical strength [17, 28, 29].
2.4.4 Human Liver Tumor Specimens for Ex Vivo Treatments
Liver tumor specimens were obtained by consent from patients with resectable
HCC, CLM, or CC tumors in accordance with the Institutional IRB approved protocol
Figure 2-2 Ex Vivo Tissue Ablation Setup. Histotripsy dose experiments were conducted using a 700kHz transducer with the focus aligned in the center of the targeted tissue, with guidance from ultrasound imaging aligned orthogonally (A-B). Tissues were imbedded within the center of a gelatin phantom (C).
33
(Carilion Clinic, Roanoke Memorial Hospital, Roanoke, VA). Patients with either biopsy-
proven or radiologically diagnosed liver tumors were consented and enrolled in the
study. At the time of resection, the resected tissues were sent fresh to pathology for
processing, and any tissue above what was needed for clinical evaluation was donated
to this study. When possible, surrounding liver parenchyma (LP) was also collected and
used to evaluate the histotripsy doses for ablating LP tissue within the surgical margin.
After harvesting, the tumor specimens were placed in a 0.9% degassed saline solution.
Before treatment, samples were degassed inside a saline solution in a vacuum chamber
(20-25 kPa) for 30-60 minutes. After degassing, the harvested tumor samples were
sectioned into 0.5-1cm3 samples and embedded in 7.5% degassed gelatin (porcine
gelatin, Millipore Sigma) tissue phantoms for treatment (Fig.2C). Gel phantoms
containing the embedded tumor samples were then refrigerated for up to 8 hours, with
all specimens treated within 24 hours of the initial harvest.
2.4.5 Histotripsy Ex Vivo Ablation Procedure
The treatment doses required to ablate HCC, CLM, and CC tumors were
investigated by histotripsy applying to the excised tissue samples embedded inside
gelatin phantoms, as described above (Fig.2). For all experiments, the 700 kHz
histotripsy therapy transducer was used to apply 5 cycle pulses to the center of the
tissue samples at a PRF of 500 Hz. These parameters closely match the histotripsy
parameters used in previous shock-scattering histotripsy liver ablation studies [15, 16,
18, 27]. For each treatment, tissue samples embedded in gelatin were mounted to a
34
motorized 3D positioning system and aligned to the focus of the therapy transducer
(Fig.2). To guide treatments and visualize the histotripsy cavitation bubble cloud in real-
time, an ultrasound imaging probe (SL18-5, AIXPLORER, SuperSonic Imagine) was
mounted perpendicularly to the tissue and aligned to the focal location of the therapy
transducer (Fig.2B). The focus of the histotripsy transducer was located and marked on
the imaging screen before each experiment by generating a bubble cloud in open water
and then marking the location on the imaging screen before aligning the tissue at the
focus. Tissue samples were then positioned such that the focal point of the transducer
was located at the desired location within the tissue.
The pressure required to generate histotripsy cavitation inside each tissue
sample was measured by gradually increasing the pressure at the focus until a
cavitation cloud was identified on US imaging. The pressure at which a consistent
histotripsy bubble cloud was generated inside each tissue sample was recorded as the
cavitation cloud threshold before conducting volumetric ablation experiments in each
sample.
Histotripsy was applied to the targeted volume at a pressure level chosen to be
~30% above the cavitation threshold, which corresponded to treatments with applied p-
between 19 MPa and 33 MPa, depending upon the sample. Volumetric treatments were
then applied automatically to a predetermined 3D grid of equally spaced treatment
points, with 2.5 mm spacing in the axial direction and 1 mm spacing in the later
directions to ensure overlap between adjacent bubble clouds. For each experiment, this
grid of points was treated with histotripsy by mechanically scanning the therapy focal
zone to cover the targeted portion of the tumor sample. The ideal treatment was a 5mm
35
x 5mm x 5mm cube consisting of 75 total points at which to dwell. Given that some
samples were too small to achieve this with a contained ablation volume, not all
ablations were this large. The desired size for each sample was 15mm x 15mm x
15mm. However, due to the irregular shapes of the tumors and the small size of some
samples, this was not always achievable. The smallest sample treated in this study was
a CC tumor that was approximately 8mm x 8mm x 8mm, which led to a treatment size
of 2.5mm x 3mm x 3mm.
For each tumor type, three to five groups were treated with histotripsy at different
doses to characterize the extent of tissue ablation generated at different histotripsy
doses (i.e., number of pulses/point). The number of pulses applied per point ranged
from 500-2000 pulses/point for HCC and CLM tumors and 1000-4000 pulses/point for
CC tumors. These treatment doses were chosen based on previous liver histotripsy
studies [15, 36, 43], with the higher doses used for CC tumors based upon a
combination of previous histotripsy studies on excised tissues [44], the relationship
between histotripsy susceptibility and mechanical properties [31, 32, 44], and literature
showing CC have an increased Young’s modulus [34]. In addition to the liver tumor
ablation experiments, a separate set of treatments was conducted on LP tissues
collected from the surgical margins around the tumors at doses of 250-1500
pulses/point. A sample size of n=3 per tissue type and treatment dose was used for all
36
experiments when possible. However, due to limitations in tissue availability, this
sample size could not be achieved for all tissues and all doses. Based on the availability
of donated tissue samples, a total of 49 experimental treatments were conducted in this
study including 8 HCC tumor samples, 15 CLM tumor samples, 12 CC tumor samples,
and 9 LP tissue samples. For each tissue, the sample numbers treated per dose is
listed in Table 1.
Table 2-1 Ex Vivo sample numbers by tissue type and Dose
2.4.6 Histology & Morphological Analysis
After treatment, tissue specimens were visually inspected and fixed in 10% formalin
for at least 24 hours before sectioning and staining. Given that each treatment was
focused on the center portion of the tissue, the central region of each tissue was
sectioned into a cassette. Using standard protocols, all tissues were paraffin embedded
and stained using hematoxylin and eosin (H&E), by ViTALS (Virginia Tech Animal
Laboratory Services, Blacksburg, VA), to assess for histotripsy damage to the tissue
parenchyma and any other changes in structure and density of collagen and other
Pulses/Point Dose
250 500 1000 1500 2000 4000
HCC n=2 n=2 n=1
CLM n=3 n=3 n=3
CC n=3 n=3 n=3 n=2
LP n=3 n=4 n=2
37
tissue structures within the ablation volume. To determine the estimated extent of tissue
ablated within the targeted volume, regions of cellular damage were compared with
regions that were outside of the ablation zone. Results were then verified by a
veterinary pathologist (SCO) and a human pathologist (DG) to ensure accuracy and
avoid biases. Histology images are representative of the samples treated.
2.5 Results
2.5.1 Histotripsy Ablation of Subcutaneous PDX CC Tumors
Well–defined histotripsy bubble clouds were observed at the focal locations in all
tumors at the beginning of the treatment (Fig.1B). Throughout the automated volumetric
treatments, the bubble clouds were continuously observed at the focus as dynamically
changing hyperechoic regions within the tissue. After treatment, histotripsy ablation
zones could be visualized on ultrasound imaging as hypoechoic regions that
corresponded with the regions exposed to the bubble cloud. This was true at each of
the three doses tested, with more clearly defined hypoechoic regions observed on
ultrasound imaging as the treatment dose increased from 250 to 1000 pulses/point
(Fig.3B-D). Histological analysis of the ablation zones showed complete ablation (no
remaining intact cells) of the CC tumors within the center of the treated region after 250
pulses/point treatments (Fig.3J). Similarly, results for the 500 pulses/point and 1000
pulses/point treatment groups showed complete ablation of the CC tumors inside the
treated regions (Fig3.K-L). The histological analysis also showed more well-defined
38
treatment margins with increasing treatment dose (Fig.3F-H). More specifically, results
from the 250 pulses/point and 500 pulses/point treatments showed small (<150µm)
regions of disrupted tissue on the margins between ablated and intact tissue that
included intact cells within the boundary region (Fig.3F-G). In contrast, these partially
treated boundary regions were not observed for the 1000 pulses/point treatments that
contained a well-defined boundary between the completely treated region and the
surrounding intact tissue (Fig.3H).
Figure 2-3 Dotted orange circles on US images (A-D) indicate approximate region of ablation. Low magnification H&E images shows ablation margins (E-H). High magnification H&E images of control and ablated region (J-L). Black boxes on 10x images indicate area magnified in respective 40x images. Scale bar on 10x H&E images = 100um (E-H), 40x scale bar = 50um (I-J).
39
2.5.2 Histotripsy Bubble Cloud Generation in Ex Vivo Tissue Specimens
The p- required to generate histotripsy bubble clouds inside each of the different
tissue types using the 700kHz clinical histotripsy transducer was measured to be
22.2±1.2 MPa, 17.7±3.1 MPa, 16.5±2.6 MPa, and 20.6+4.6 for HCC, CLM, CC, and LP
samples, respectively. In general, the bubble clouds in all tissue types showed typical
histotripsy features with a dense dynamically changing hyperechoic bubble cloud
observed within or near the focal region of the transducer, as visualized on US imaging.
During volumetric ablation experiments, the hyperechoic bubble clouds were maintained
throughout the treatments in all tissue experiments. In some samples, particularly
samples smaller than the desired 15mm x 15 mm x 15 mm dimensions, the cavitation
bubble clouds formed at the interface of the gelatin phantom and the tissue including
instances of both prefocal and post-focal cavitation clouds. These effects were
observed more often in the CLM and CC samples. In cases in which prefocal cavitation
was noted during the initial bubble cloud generation, the treatment zone was adjusted to
a deeper location in the sample when possible and the volumetric treatments were only
included as part of this study for samples in which a focal bubble cloud remained
present for the duration of the treatment.
2.5.3 Histotripsy Ablation of Ex Vivo HCC Specimens
For HCC samples treated with 500 or more pulses/point (Fig.4B-D/F-H), histotripsy
resulted in complete ablation of the targeted tumor volume with no notable regions of
intact cells remaining inside the ablated area. There was tumor necrosis characterized
40
by remaining cell nuclei and other subcellular tissue debris inside the treated regions on
these samples, with the number of these nuclei reduced in concentration for the higher
treatment doses. At 1000 pulses/point or more, there was a complete loss of cellular
structures characterized as tissue ablated to amorphous debris.
2.5.4 Histotripsy Ablation of Ex Vivo CLM Specimens
Histological analysis demonstrated partial tumor ablation for CLM samples treated
with 500 pulses/point, with more than 75% of cells within the treated region remaining
after treatment (Fig.5B/G). Small, sporadic foci of ablation were observed inside of
these samples, with the majority of the treatment volume remaining intact. For 1000
pulses/point CLM samples, a larger portion of the tumor samples were observed to be
ablated after treatment with ~25-75% of the targeted region containing viable tumor
Figure 2-4 Comparison of untreated human HCC tumors with ex vivo ablations at 500 pulses/point, 1000 pulses/point, and 2000 pulses/point. 10x magnification scale bar= 100um (A-D), 40x scale bar= 50um (E-H). Black boxes on 10x images indicate area magnified in respective 40x images. Histology images are representative of the samples treated.
41
tissue after histotripsy (Fig.5C/H). At the higher treatment doses of 1500 and 2000
pulses/point, histotripsy was observed to generate significantly more tissue damage
with complete ablation of the targeted tumor regions observed in some of the samples
at both doses (Fig.5D-E/I-J). However, results showed a large amount of variability in
the extent of ablation among treatments within each group, with some samples showing
only partial tumor ablation. More specifically, the number of viable cells remaining after
treatment ranged from <5% (complete ablation) to ~25% in each group, with complete
ablation achieved in 2 of the 3 samples at 1500 pulses/point and 1 of the 2 samples
tested at 2000 pulses/point. It was noted that the tissues that did not show complete
Figure 2-5 Comparison of untreated human CLM tumors with those treated at 500 pulses/point, 1000 pulses/point, 1500 pulses/point, and 2000 pulses/point. 10x magnification scale bar= 100um (A-E), 40x scale bar= 50um (F-J). Black boxes on 10x images indicate area magnified in respective 40x image. Histology images are representative of the samples treated.
42
ablation within these groups appeared to contain more glandular structures, thus, were
more well-differentiated. Less differentiated tumors with poor cellular organization had
greater ablation. Similarly, histological analysis showed the CLM tumors tested in this
study exhibited varying degrees of fibrosis within the tumor. Partial ablations were most
often exhibited by CLM tumors with a robust, desmoplastic (or fibrous) response. For
example, for the 1000 pulses/point samples shown in Figure 5, the ablated regions of
the tumor are found sporadically within the more fibrous regions of the tissue showing
loss of cellular detail but still maintaining the fibrotic tissue architecture (Fig.5C/H). In
contrast, the extent of ablation was significantly higher within the less fibrotic regions of
these tumors. This effect can be seen in the example 2000 pulses/point image shown in
Figure 5, in which large regions of the tissue were fully ablated, with the remaining
Figure 2-6 Comparison of untreated human CC tumors with those treated at 1000 pulses/point, 1500 pulses/point, 2000 pulses/point, and 4000 pulses/point. 10x magnification scale bar= 100um (A-E), 40x scale bar= 50um (F-J). Black boxes on 10x images indicate area magnified in respective 40x image. Histology images are representative of the samples treated.
43
tissue lacking cellular detail but with a few nuclei corresponding to the more fibrotic
portions of the tumor.
2.5.5 Histotripsy Ablation of Ex Vivo CC Specimens
Histological analysis demonstrated partial tumor ablation for CC samples treated
with 1000 pulses/point, with sporadic foci of ablated tumor observed throughout the
treated volume with interspersed regions of intact tumor (Fig.6B/G). Results for the
1500 pulses/point and 2000 pulses/point treatments showed similar results with larger
regions of complete loss of cellular detail, supporting a completely ablated portion of the
tumor in these samples (Fig.6) compared to the 1000 pulses/point group (Fig.6B/G).
For example, the 2000 pulses/point sample shown in Figure 6 has two visible regions of
tumor that were not fully ablated, both of which correspond to more fibrotic regions of
the tissue. In contrast, the fully ablated regions within this sample appear to correspond
to less dense and less fibrotic regions of the tumor. At the highest treatment dose of
4000 pulses/point, histotripsy again resulted in a similar result with only partial ablation
generating within each of the CC tumors and ~50-75% of the cells showing viable nuclei
but with loss of cytoplasmic detail within the ablation region remaining after treatment
(Fig.6E/J). In these samples, the sporadic foci of ablation again appeared to be located
in the regions of the tissue low in fibrosis (Fig.6). Looking at these foci under higher
magnification, the tissue within these ablated regions showed complete ablation with no
remaining viable cells. In contrast, the less ablated regions of the CC tumors were
highly more fibrotic with >75-90% of viable cells remaining after treatment, as shown in
44
Supplemental Figure S1. Finally, it is important to note the high amount of variability in
the extent of ablation was observed among treatments within each group. Several of the
treatments resulted in >75% ablation (<25% of viable cells remaining) at dosing
regimens of 1000, 1500, 2000, and 4000 pulses/point whereas other treatments
resulted in <25% ablation with the majority of the targeted region remaining intact after
treatment.
2.5.6 Histotripsy Ablation of Ex Vivo LP Specimens
Part of the surrounding liver parenchyma was excised in addition to the tumor for six
patients enrolled in this study. Four patients had hepatic steatosis as identified by
abdominal imaging. One patient had hemochromatosis in addition to steatosis, one was
Hepatitis C positive with fibrosis, and another patient had no identifiable liver disease.
These samples were divided based on the presence of steatosis. On histology, tissues
Figure 2-7 Supplemental Figure S1. Single sample of Cholangiocarcinoma Ablated at 4000 pulses/point illustrating fibrosis dependence to susceptibility. Region of fibrosis (A) showed no damage, while less fibrotic region (B) had sporadic ablated foci. High magnification of the region in B indicated by the black box is Figure 7E 4000 pulses/point location. Scale bars = 500um. * Indicate ablated region.
45
from patients with hepatic steatosis were readily apparent based on the presence of
macrovesicular fat globules within the tissues. For non-steatotic samples, a sample size
of n=1 was conducted at 250, 500, and 1000 pulses/point. Steatotic samples were
treated at n=2 at 250 pulses/point, n=3 for 500 pulses/point, and n=1 for 1000
pulses/point. A complete ablation was formed within the tissues that did not have
hepatic steatosis even at doses as low as 250 pulses/point (Fig.7). In comparison,
tissues with hepatic steatosis were noted to have sporadic foci of ablations at 250
pulses/point with more complete ablation generated at the higher doses. For instance,
only a few remaining viable cells were visible inside the ablation zone after 500
pulses/point and complete ablation of the treated region was observed after 1000
pulses/point (Fig.8).
Figure 2-8 Non-steatotic liver parenchyma treatment at 250 pulses/point, 500 pulses/point, and 1000 pulses/point demonstrating less uniform ablations. 10x magnification scale bar= 100um (A-D), 40x scale bar= 50um (E-H). Black boxes on 10x images indicate area magnified in respective 40x images. Histology images are representative of the samples treated.
46
2.6 Discussion
To study the potential of histotripsy for CC tumor ablation, this work first investigated
the in vivo feasibility of histotripsy CC ablation in a PDX mouse model, with results
showing histotripsy could achieve precise and complete ablation within the targeted
region of the tumor. The treatment doses used to treat the CC tumors in this study were
higher than those reported previously for xenografted HCC tumors [28]. Given the
known correlation between tissue mechanical properties and susceptibility to histotripsy,
we hypothesize that this difference is likely due to the higher mechanical strength of CC
tumors [31-33]. Overall, the results from this in vivo feasibility study suggest that
histotripsy will be capable of treating CC tumors if the proper treatment dose is applied.
In the second part of this study, the effectiveness of histotripsy for ablating CC tumors
was compared to HCC and CLM tumors, as well as LP tissue from the surgical margin,
Figure 2-9 Steatotic liver parenchyma treatment at 250 pulses/point, 500 pulses/point, and 1000 pulses/point demonstrating less uniform ablations. 10x magnification scale bar= 100um (A-D), 40x scale bar= 50um (E-H). Black boxes on 10x images indicate area magnified in respective 40x images. Arrows indicating example cells within an ablated region that appear to be intact. Histology images are representative of the samples treated.
47
in ex vivo experiments using human tumor specimens. Overall, the results of these
experiments help to demonstrate the histotripsy dosimetry metrics for treating CC in
comparison to HCC and CLM liver tumors. More specifically, the results showed that
both the CC and CLM tumors required higher treatment doses before tissue ablation
was observed in comparison to the softer HCC and LP samples that showed complete
tumor ablation at lower treatment doses. These findings match prior studies that have
shown fibrous tissues, which have been reported to have higher mechanical strength,
are more resistant to histotripsy and therefore require higher treatment doses [31-33].
Results from HCC treatments demonstrated that HCC can be completely ablated with
histotripsy with less than 500 pulses/point, agreeing with previous reports that
histotripsy has the potential for rapid, precise, and complete ablation of primary liver
tumors. Since CLM tumors are stiffer than HCC tumors [31, 32], we hypothesized that
higher treatment doses would be required for generating ablation in these samples.
Results supported this hypothesis showing histotripsy was able to achieve only small
regions of partial ablation for 500 pulses/point. At higher treatment doses of 1500 and
2000 pulses/point, nearly complete or complete ablation was observed in several CLM
samples, with the only untreated regions corresponding to highly fibrous regions within
the tumor sample.
For ex vivo experiments with CC tumors, the trends were similar to those
observed for CLM with higher doses again being needed to generate ablation in the CC
tumors. Even at the highest doses treated in this study (4000 pulses/point), there were
no cases in which we observed complete ablation of the entire treatment volume within
the CC tumors, suggesting that higher doses will be needed to achieve full ablation of
48
these highly fibrous CC tumors. This finding appeared to be at least partially due to the
large variation in tissue composition between samples as well as the heterogeneity of
the tissue within a given sample (Fig.S1). Partial ablations were more pronounced in
the CC and CLM tumor samples. We hypothesize that this is due to the higher
mechanical strength and the more fibrous nature of these tumors.
Although complete ablation was not generated in the ex vivo CC tumors,
significant ablation was observed for treatments at the two highest doses tested, with
nearly 75% of the treated volume ablated after 2000 and 4000 pulses/point. Overall,
these results highlight the need for tumor-specific treatment strategies in future clinical
treatments of liver tumors with histotripsy to ensure complete ablation of CLM and CC
tumors and more efficient ablation of HCC tumors. For instance, these approaches
could include the development of guidelines for the treatment doses (histotripsy
pulses/point) needed for treating different tumor types as well as the development of
optimized pulsing parameters, although further work would be needed in order to
investigate this possibility. Additionally, improved real-time feedback methods could be
utilized to ensure complete therapeutic ablation of specific tumors such as those
previously developed using bubble-induced color doppler feedback [45, 46] or cavitation
detection [47].
In addition to the differences in ablation observed between tumor types, another
important observation from this study was the variability in ablation between samples of
a given tumor type. This result was possibly due in part to variations in the tissue
composition, mainly fibrotic deposits, between any two tumors of the same type as well
as the heterogeneity of the tissue within a single tumor. This effect would explain why
49
the more uniform and softer HCC samples consistently resulted in complete ablation at
lower treatment doses compared to more variable results and less complete ablation
observed in the more heterogeneous and fibrous CLM and CC tumors. Looking at the
regions of partial ablation in these samples, results showed that the foci of ablation
interspersed within regions that weren’t ablated corresponded to the areas with less
fibrosis. This result suggests that there is a need for ensuring histotripsy treatment
doses are either optimized for each patient based on pretreatment assessments of the
tumor composition and mechanical character, such as through MRI or ultrasound
elastography. Alternatively, a sufficient treatment dose could be identified based on the
most fibrous tumors within each type based upon historically determined mechanical
properties. For example, it is possible that the different subtypes of intrahepatic CC
(mass-forming, periductal, intraductal) may respond differently to histotripsy both in the
required treatment dose as well as the efficacy of histotripsy for achieving a desired
clinical outcome. Future work should explore this possibility in order to establish the
optimal role of histotripsy for the treatment of each subtype of intrahepatic CC, as well
as extrahepatic CC, and determine if tumor type specific treatment methods are
required.
However, although we believe the variations in tissue composition are likely to
explain some of the variability in our ex vivo tissue experiments and should be
considered in future clinical histotripsy treatments, it is also important to note that these
findings could also be due to the limitations of our tissue collection methods in this
study. Unfortunately, due to the requirements of the clinical pathologists before tissue
collection, the samples collected for this study were often smaller than the desired size
50
for performing optimal ex vivo histotripsy experiments. In these smaller samples,
prefocal bubble clouds were more often observed on the tissue-gel interface especially
in the more fibrous CLM and CC tumors, preventing a more uniform histotripsy
treatment from being applied to the tissue volume and likely contributing to some of the
variable ablation results observed in those tissues. These effects are not expected to
occur in vivo where much larger tumors that are surrounded by adjacent LP will be
treated with histotripsy, therefore minimizing the risk of prefocal cavitation that causes
inconsistent tumor ablation.
Another observation from the ex vivo experiments conducted in this work was the
finding that non-steatotic LP was readily ablated with histotripsy, similar to HCC
samples, whereas higher treatment doses were required to ablate steatotic LP samples.
This finding is important for future liver cancer treatments with histotripsy since it is a
standard clinical practice to ablate a margin of the surrounding liver around the tumor to
destroy all microscopic foci of tumor cells that might be within these regions. The finding
that the LP surrounding tumors in patients with hepatic steatosis, or fatty liver, required
higher treatment doses compared to those without this disease, should be noted in
future studies optimizing treatment parameters for hepatic histotripsy. It is also possible
that fatty pockets dispersed throughout the liver could contribute to more diffuse or
prefocal cavitation. This effect would make sense since previous work has shown a
significantly lower cavitation threshold for fat compared to water-based parenchymal
tissues [20, 21]. However, since no consistent differences in cavitation were observed
between the steatotic and non-steatotic LP samples treated in this study, further work
remains needed to investigate the role that fat deposits play in hepatic histotripsy and
51
whether any differences in the cavitation threshold can cause variation in ablation
completeness similar to what has been observed for differences in tissue fibrosis.
2.7 Conclusion
This study tested the potential of histotripsy for the treatment of CC tumors for the
first time, with results suggesting that histotripsy has the potential to be used as a non-
invasive ablation method for the treatment of CC. Results from the in vivo experiments
demonstrated that histotripsy could generate precise and complete ablation of CC
tumors in a patient-derived xenograft CC mouse model. In the second part of this work,
which compared the histotripsy treatment dose required to ablate different types of liver
tumors, results showed that CC and CLM tumors required significantly higher doses
than HCC tumors. These findings highlight the need for tumor-specific treatment
strategies for future histotripsy liver cancer procedures to ensure more fibrous tumor
types, as well as more fibrous regions within a tumor, are completely ablated during
histotripsy. Overall, the findings of this study support further investigation of histotripsy
for the treatment of CC in addition to HCC and CLM.
2.8 Acknowledgement
This research was supported by a grant from the Focused Ultrasound
Foundation (FUSF-RAP-S-18-00011). The authors would also like to thank the ICTAS
Center for Engineered Health, the Virginia Tech Department of Biomedical Engineering
52
and Mechanics, Virginia Tech Carilion School of Medicine, and the Virginia-Maryland
College of Veterinary Medicine for their support of this study. This work was conducted
in memory of Andrew J. Lockhart, who passed away from cholangiocarcinoma on
September 30, 2016. The authors encourage anyone interested to consider a donation
to the Focused Ultrasound Foundation in Andrew’s name.
53
2.9 References
[1] ACS, "Cancer Facts & Figures 2020," American Cancer Society, 2020.
[2] M. C. Jansen, R. van Hillegersberg, R. A. Chamuleau, O. M. van Delden, D. J.
Gouma, and T. M. van Gulik, "Outcome of regional and local ablative therapies
for hepatocellular carcinoma: a collective review," Eur J Surg Oncol, vol. 31, no.
4, pp. 331-47, May 2005, doi: 10.1016/j.ejso.2004.10.011.
[3] L. Rossi et al., "Current approach in the treatment of hepatocellular carcinoma,"
World J Gastrointest Oncol, vol. 2, no. 9, pp. 348-59, Sep 15 2010, doi:
10.4251/wjgo.v2.i9.348.
[4] D. M. Nagorney, J. H. Donohue, M. B. Farnell, C. D. Schleck, and D. M. Ilstrup,
"Outcomes after curative resections of cholangiocarcinoma," Archives of surgery,
vol. 128, no. 8, pp. 871-878, 1993.
[5] K. Feng et al., "A randomized controlled trial of radiofrequency ablation and
surgical resection in the treatment of small hepatocellular carcinoma," Journal of
hepatology, vol. 57, no. 4, pp. 794-802, Oct 2012, doi:
10.1016/j.jhep.2012.05.007.
[6] M. S. Chen et al., "A prospective randomized trial comparing percutaneous local
ablative therapy and partial hepatectomy for small hepatocellular carcinoma," (in
eng), Ann Surg, Randomized Controlled Trial, Research Support, Non-U.S. Gov't
vol. 243, no. 3, pp. 321-8, Mar 2006, doi: 10.1097/01.sla.0000201480.65519.b8.
54
[7] C. W. Hammill, K. G. Billingsley, M. A. Cassera, R. F. Wolf, M. B. Ujiki, and P. D.
Hansen, "Outcome after laparoscopic radiofrequency ablation of technically
resectable colorectal liver metastases," (in eng), Annals of surgical oncology, vol.
18, no. 7, pp. 1947-54, Jul 2011, doi: 10.1245/s10434-010-1535-9.
[8] G. Otto, C. Duber, M. Hoppe-Lotichius, J. Konig, M. Heise, and M. B. Pitton,
"Radiofrequency ablation as first-line treatment in patients with early colorectal
liver metastases amenable to surgery," Ann Surg, vol. 251, no. 5, pp. 796-803,
May 2010, doi: 10.1097/SLA.0b013e3181bc9fae.
[9] R. Salem et al., "Institutional Decision to Adopt Y90 as Primary Treatment for
HCC Informed by a 1,000-patient 15-year Experience," Hepatology, Dec 1 2017,
doi: 10.1002/hep.29691.
[10] A. Forner, J. M. Llovet, and J. Bruix, "Hepatocellular carcinoma," Lancet, vol.
379, no. 9822, pp. 1245-55, Mar 31 2012, doi: 10.1016/S0140-6736(11)61347-0.
[11] L. European Association for the Study of the, R. European Organisation for, and
C. Treatment of, "EASL–EORTC Clinical Practice Guidelines: Management of
hepatocellular carcinoma," Journal of hepatology, vol. 56, no. 4, pp. 908-943,
2012. [Online]. Available:
http://linkinghub.elsevier.com/retrieve/pii/S0168827811008737?showall=true.
[12] A. Gillams et al., "Thermal ablation of colorectal liver metastases: a position
paper by an international panel of ablation experts, The Interventional Oncology
Sans Frontieres meeting 2013," Eur Radiol, vol. 25, no. 12, pp. 3438-54, Dec
2015, doi: 10.1007/s00330-015-3779-z.
55
[13] N. Rozenblum et al., "Oncogenesis: An "Off-Target" Effect of Radiofrequency
Ablation," Radiology, vol. 276, no. 2, pp. 426-32, Aug 2015, doi:
10.1148/radiol.2015141695.
[14] B. Sangro et al., "Prevention and treatment of complications of selective internal
radiation therapy: Expert guidance and systematic review," Hepatology, vol. 66,
no. 3, pp. 969-982, Sep 2017, doi: 10.1002/hep.29207.
[15] C. M. Smolock AR, Vlaisavljevich E, Gendron-Fitzpatrick A, Green C, Ziemlewicz
TJ and Lee FT, "Robotically Assisted Sonic Therapy as a Non-Invasive Non-
Thermal Ablation Modality: Proof of Concept in a Porcine Liver Model,"
Radiology, vol. 287, no. 2, pp. 485-93, 2018.
[16] S. X. Vidal Jove J, Vlaisavljevich E, Cannata J, Duryea A, Miller R, Lee F and
Ziemlewicz T, "Phase I Study of Safety and Efficacy of Hepatic Histotripsy:
Preliminary Results of First in Man Experience with Robotically-Assisted Sonic
Therapy," in International Society of Therapeutic Ultrasound, Barcelona, Spain,
2019.
[17] E. Vlaisavljevich et al., "Non-Invasive Ultrasound Liver Ablation Using Histotripsy:
Chronic Study in an In Vivo Rodent Model," Ultrasound Med Biol, vol. 42, no. 8,
pp. 1890-902, Aug 2016, doi: 10.1016/j.ultrasmedbio.2016.03.018.
[18] E. Vlaisavljevich et al., "Image-guided non-invasive ultrasound liver ablation
using histotripsy: feasibility study in an in vivo porcine model," Ultrasound Med
Biol, vol. 39, no. 8, pp. 1398-409, Aug 2013, doi:
10.1016/j.ultrasmedbio.2013.02.005.
56
[19] T. Worlikar et al., "Histotripsy for Non-Invasive Ablation of Hepatocellular
Carcinoma (HCC) Tumor in a Subcutaneous Xenograft Murine Model," Conf Proc
IEEE Eng Med Biol Soc, vol. 2018, pp. 6064-6067, Jul 2018, doi:
10.1109/EMBC.2018.8513650.
[20] A. D. Maxwell, C. A. Cain, T. L. Hall, J. B. Fowlkes, and Z. Xu, "Probability of
cavitation for single ultrasound pulses applied to tissues and tissue-mimicking
materials," Ultrasound Med Biol, vol. 39, no. 3, pp. 449-65, Mar 2013, doi:
10.1016/j.ultrasmedbio.2012.09.004.
[21] E. Vlaisavljevich et al., "Effects of ultrasound frequency and tissue stiffness on
the histotripsy intrinsic threshold for cavitation," Ultrasound Med Biol, vol. 41, no.
6, pp. 1651-67, Jun 2015, doi: 10.1016/j.ultrasmedbio.2015.01.028.
[22] Z. Xu et al., "Controlled ultrasound tissue erosion," IEEE Trans Ultrason
Ferroelectr Freq Control, vol. 51, no. 6, pp. 726-36, Jun 2004, doi:
10.1109/tuffc.2004.1308731.
[23] R. M. Xu Z, Hall TL, Mycek M-A, Fowlkes JB and Cain CA, "Evolution of bubble
clouds produced in pulsed cavitational ultrasound therapy - histotripsy," IEEE
Trans Ultrason Ferroelectr Freq Control, vol. 55, pp. 1122-32, 2008.
[24] E. Vlaisavljevich, A. Maxwell, M. Warnez, E. Johnsen, C. A. Cain, and Z. Xu,
"Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
mechanical properties," IEEE Trans Ultrason Ferroelectr Freq Control, vol. 61,
no. 2, pp. 341-52, Feb 2014, doi: 10.1109/TUFFC.2014.6722618.
[25] E. Vlaisavljevich, A. Maxwell, L. Mancia, E. Johnsen, C. Cain, and Z. Xu,
"Visualizing the Histotripsy Process: Bubble Cloud-Cancer Cell Interactions in a
57
Tissue-Mimicking Environment," Ultrasound Med Biol, vol. 42, no. 10, pp. 2466-
77, Oct 2016, doi: 10.1016/j.ultrasmedbio.2016.05.018.
[26] V. E. Kim Y, Owens GE, Allen SP, Cain CA and Xu Z, "In vivo transcostal
histotripsy therapy without aberration correction," Phys Med Biol, vol. 59, pp.
2553-68, 2014.
[27] E. Vlaisavljevich et al., "Non-Invasive Liver Ablation Using Histotripsy: Preclinical
Safety Study in an In Vivo Porcine Model," Ultrasound Med Biol, vol. 43, no. 6,
pp. 1237-1251, Jun 2017, doi: 10.1016/j.ultrasmedbio.2017.01.016.
[28] T. Worlikar et al., "Histotripsy for Non-Invasive Ablation of Hepatocellular
Carcinoma (HCC) Tumor in a Subcutaneous Xenograft Murine Model," in 2018
40th Annual International Conference of the IEEE Engineering in Medicine and
Biology Society (EMBC), 2018: IEEE, pp. 6064-6067.
[29] S. Qu et al., "Non-thermal histotripsy tumor ablation promotes abscopal immune
responses that enhance cancer immunotherapy," Journal for ImmunoTherapy of
Cancer, vol. 8, no. 1, 2020.
[30] S. K. Venkatesh et al., "MR elastography of liver tumors: preliminary results,"
American Journal of Roentgenology, vol. 190, no. 6, pp. 1534-1540, 2008.
[31] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," (in eng), Phys Med Biol, vol. 59, no. 2, pp. 253-70, Jan 20 2014, doi:
10.1088/0031-9155/59/2/253.
[32] E. Vlaisavljevich, A. Maxwell, M. Warnez, E. Johnsen, C. Cain, and Z. Xu,
"Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
58
mechanical properties," (in eng), IEEE Transactions on Ultrasonics,
Ferroelectrics, and Frequency Control, vol. 61, no. 2, pp. 341-52, Feb 2014, doi:
10.1109/TUFFC.2014.6722618.
[33] E. Vlaisavljevich, Z. Xu, A. Arvidson, L. Jin, W. Roberts, and C. Cain, "Effects of
thermal preconditioning on tissue susceptibility to histotripsy," Ultrasound in
medicine & biology, vol. 41, no. 11, pp. 2938-2954, 2015.
[34] A. Guibal et al., "Evaluation of shearwave elastography for the characterisation of
focal liver lesions on ultrasound," European radiology, vol. 23, no. 4, pp. 1138-
1149, 2013.
[35] T. G. Schuster, J. T. Wei, K. Hendlin, R. Jahnke, and W. W. Roberts, "Histotripsy
Treatment of Benign Prostatic Enlargement Using the Vortx Rx System: Initial
Human Safety and Efficacy Outcomes," Urology, vol. 114, pp. 184-187, Apr
2018, doi: 10.1016/j.urology.2017.12.033.
[36] E. Vlaisavljevich et al., "Non-invasive liver ablation using histotripsy: preclinical
safety study in an in vivo porcine model," Ultrasound in medicine & biology, vol.
43, no. 6, pp. 1237-1251, 2017.
[37] W. W. Roberts, D. Teofilovic, R. C. Jahnke, J. Patri, J. M. Risdahl, and J. A.
Bertolina, "Histotripsy of the prostate using a commercial system in a canine
model," The Journal of urology, vol. 191, no. 3, pp. 860-865, 2014.
[38] A. D. Maxwell et al., "Cavitation clouds created by shock scattering from bubbles
during histotripsy," J Acoust Soc Am, vol. 130, no. 4, pp. 1888-98, Oct 2011, doi:
10.1121/1.3625239.
59
[39] J. E. Parsons, C. A. Cain, G. D. Abrams, and J. B. Fowlkes, "Pulsed cavitational
ultrasound therapy for controlled tissue homogenization," (in eng), Ultrasound in
Medicine and Biology, vol. 32, no. 1, pp. 115-29, Jan 2006, doi: S0301-
5629(05)00374-1 [pii]
10.1016/j.ultrasmedbio.2005.09.005.
[40] J. E. Parsons, C. A. Cain, and J. B. Fowlkes, "Cost-effective assembly of a basic
fiber-optic hydrophone for measurement of high-amplitude therapeutic ultrasound
fields," J Acoust Soc Am, vol. 119, no. 3, pp. 1432-40, Mar 2006, doi:
10.1121/1.2166708.
[41] E. Vlaisavljevich, T. Gerhardson, T. Hall, and Z. Xu, "Effects of f-number on the
histotripsy intrinsic threshold and cavitation bubble cloud behavior," Phys Med
Biol, vol. 62, no. 4, pp. 1269-1290, Feb 21 2017, doi: 10.1088/1361-
6560/aa54c7.
[42] R. M. Brock et al., "Patient derived xenografts expand human primary pancreatic
tumor tissue availability for ex vivo irreversible electroporation testing," Frontiers
in Oncology, vol. 10, 2020.
[43] K. C. Longo et al., "Histotripsy Ablations in a Porcine Liver Model: Feasibility of
Respiratory Motion Compensation by Alteration of the Ablation Zone Prescription
Shape," CardioVascular and Interventional Radiology, pp. 1-7, 2020.
[44] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," Physics in Medicine & Biology, vol. 59, no. 2, p. 253, 2013.
60
[45] J. J. Macoskey et al., "Bubble-induced color Doppler feedback correlates with
histotripsy-induced destruction of structural components in liver tissue,"
Ultrasound in medicine & biology, vol. 44, no. 3, pp. 602-612, 2018.
[46] T. Y. Wang, T. L. Hall, Z. Xu, J. B. Fowlkes, and C. A. Cain, "Imaging feedback of
histotripsy treatments using ultrasound shear wave elastography," (in eng), IEEE
Transactions on Ultrasonics, Ferroelectrics, and Frequency Control, Research
Support, N.I.H., Extramural vol. 59, no. 6, pp. 1167-81, Jun 2012, doi:
http://dx.doi.org/10.1109/TUFFC.2012.2307.
[47] J. J. Macoskey et al., "Using the cavitation collapse time to indicate the extent of
histotripsy-induced tissue fractionation," (in eng), Phys Med Biol, vol. 63, no. 5, p.
055013, Mar 8 2018, doi: 10.1088/1361-6560/aaae3b.
61
Chapter 3. Histotripsy for Non-Invasive Ablation of Cholangiocarcinoma Carcinoma in a Patient-Derived Xenograft Model
Alissa Hendricks-Wenger, Sofie Saunier, Alexander Simon, Sheryl Coutermarsh-Ott1, Irving C. Allen, Eli Vlaisavljevich
This chapter is excerpted from a manuscript in preparation for submission to Ultrasound
in Medicine and Biology.
62
3.1 Abstract
Histotripsy is a non-invasive focused ultrasound ablation method that is currently
being developed for the treatment of primary and metastatic liver tumors. In a recent
study, we investigated the feasibility of using histotripsy for the ablation of
cholangiocarcinoma (CC), a rare form of liver cancer of the bile duct that is difficult to
treat with current therapies due to its common location near critical structures, late
diagnosis, and fibrous tissue microenvironment. As part of this prior study, the in vivo
feasibility of treating CC tumors was show in an acute patient-derived xenografted
(PDX) CC mouse model. Here, we investigate histotripsy for the ablation of CC tumors
in vivo in a chronic PDX mouse model of CC. Patient-derived xenografted CC mice
were treated with histotripsy using a 1MHz transducer at a PRF of 500 Hz and an
applied dose of 500 pulses/point. First, a pilot study was conducted to determine if
histotripsy could safely treat the entire CC tumors plus a ~2mm surgical margin in the
PDX model without inducing significant adverse events. Results showed histotripsy
generated significant injuries to intestinal tissues located within the treatment margin,
precluding the use of this positive-margin treatment strategy in subsequent
experiments. For the next set of experiments, histotripsy was applied to CC tumors
(n=6) with the bubble cloud contained only to the targeted tumor (zero-margin), resulting
in ~80%-90% of the tumor treated and no cavitation events outside of the tumor. After
treatment, ablation was visualized using ultrasound imaging immediately post-treatment
and subjects were then monitored over time to determine changes in tumor size, long-
63
term survival, and safety outcomes assessed by adverse events. Results were
compared to control mice that did not receive histotripsy (n=6) and showed histotripsy
achieved an average of 73% reduction of tumor diameter 26 days after treatment, with
no significant adverse events or damage to surrounding tissues. Notably, 3 of the 6
treated tumors were reduced in diameter to an undetectable size after ~2.5 weeks as
the ablated homogenate was reabsorbed, before tumor growth was again observed.
The treated animals were found to have significantly increased progression-free and
overall survival, and no metastases were observed in any animals. Overall, results
support the hypothesis that histotripsy can be used as a safe and effect method for
treating CC tumors while also highlighting the limitations of the subcutaneous murine
model.
3.2 Keywords
Focused Ultrasound, Patient Derived Xenograft, Cholangiocarcinoma, Tumor Ablation,
Histotripsy
64
3.3 Introduction
Cholangiocarcinoma (CC) is a rare form of liver cancer that affects ~8,000 people in
the United States each year and has a 5-year relative survival rate of only 8% [1, 2].
Current treatment options include surgery, chemoembolization, and radiotherapy;
however, due to their location near vital structures, CC tumors pose a unique challenge
for these modalities [3-5]. For example, although curative resection offers the best
chance for long-term survival, only a fraction of CC tumors are considered to be
resectable, including 15% of intrahepatic CC tumors and 56% of extrahepatic CC
tumors [6]. There has been some success in using thermal ablation methods for the
treatment of the primary liver tumors such as hepatocellular cholangiocarcinoma as well
as metastatic liver tumors. However, the lack of precision and high complication rates
when in proximity to critical structures present significant limitations for thermal ablation
[7-14]. Due to this limitation, thermal ablation is often ruled out as a treatment option for
CC due to the high risk of complications for treating CC tumors which are most often
located in close proximity to the bile ducts [14].
Histotripsy is an image-guided, non-invasive, non-thermal, and non-ionizing tumor
ablation method that uses focused ultrasound to ablate targeted tissue through
controlled generation of acoustic cavitation [15-17]. Histotripsy delivers focused,
microsecond long, high-pressure ultrasound pulses that generate a cavitation “bubble
cloud” inside of the target tissue at the focus of the transducer [18-22]. The generation
of these cavitation bubbles, which rapidly expand and collapse, induces high strain to
65
cells within the tumor and results in precise, non-thermal tissue ablation [23]. Histotripsy
has shown great promise for the treatment of liver cancer in both preclinical studies [15,
16, 24-28] and early clinical trials [29]. These studies have shown that histotripsy can
safely and non-invasively ablate tumors in the liver with high precision, including regions
located near critical structures such as bile ducts, nerves, and blood vessels [22, 30-36].
More specifically, due to its non-thermal nature, histotripsy has shown the ability to
effectively ablate tissue surrounding these critical structures without causing damage to
their anatomy, likely due to the higher mechanical strength and fibrotic composition of
these structures [16, 22, 27, 31, 35].
Due to these previous studies showing the potential of hepatic histotripsy, our team
recently completed a feasibility study to investigate whether histotripsy could be used
for the ablation of CC tumors [37]. The ex vivo portion of this study demonstrated that
CC tumors require a higher dose (i.e. more histotripsy pulses per treatment point) to
reach comparable ablation qualities to other liver malignancies, specifically
hepatocellular carcinoma and colorectal carcinoma tumors that metastasized to the
liver. The higher treatment dose required for CC tumors was hypothesized to be due to
the higher mechanical strength and more fibrotic composition of the CC tumors in
comparison to hepatocellular carcinoma and metastatic liver tumors [37, 38]. This
previous study also investigated the in vivo feasibility of histotripsy for treating CC
tumors in a patient-derived xenograft (PDX) CC tumor model, with results showing
histotripsy could achieve ablation of CC tumors using a 1MHz small animal histotripsy
transducer [17, 37]. Here, we build on this prior work in order to investigate the safety
and efficacy of histotripsy for treating CC tumors in the subcutaneous PDX mouse
66
model of CC. This model was chosen due the tumor-specific tumor microenvironment
that will allow for proper assessment of tumor ablation success [39] as well as the lack
of other well-established orthotopic murine or large animal CC tumor models. In the first
part of this study, we investigated whether histotripsy could be used to safely generate
complete tumor ablation plus a positive surgical margin in the PDX tumor model. Based
on the results from these pilot studies, a second set of experiments was conducted in
order to investigate the long-term response of CC tumors treated with histotripsy using
more conservative treatments in which the histotripsy bubble cloud was contained
inside the tumor, with these zero-margin treatments resulting in ~80-90% of the tumor
volume targeted with histotripsy in order to avoid off target damage to surrounding
tissues. The acute and chronic response of the murine PDX tumors were evaluated to
determine the safety and feasibility of treating CC tumors with histotripsy through
imaging, heath monitoring, and histological analysis.
3.4 Materials and Methods
3.4.1 Tumor Injections and Health Monitoring
Seventeen female NSG mice with subcutaneous flank patient derived
xenografted (PDX) CC tumors were ordered from and prepared by Jackson Laboratory
(Bar Harbor, Maine, USA). Through a process previously described for pancreatic PDX
tumors [39], a human CC tumor was collected from a patient and propagated as a
heterogeneous tumor in immunocompromised NSG mice. The PDX tumor (TM01225)
67
was a diagnosed adenosquamous carcinoma, a subtype of CC, with an unspecified
grade/stage that was surgically resected from the bile duct, the primary site, of a 79-
year-old white/Hispanic male. Mice were delivered to our facility after confirmed tumor
engraftment. After arrival, animals were monitored for general health, weight, and tumor
size at least 3 times per week until tumors reached the desired treatment size of ~0.4cm
in diameter. Tumor diameters were measured with calipers and calculated as the
square root of two perpendicular measurements, as previously described [39]. All
animal experiments were approved and carried out in accordance with the Virginia Tech
Institutional Animal Care and Use Committee under IACUC protocol 18-031-CVM.
3.4.2 Histotripsy Setup
The in vivo histotripsy treatment was applied using a custom 8-element, 1 MHz
small animal histotripsy transducer with a geometric focus of 36 mm, an aperture size of
52.7 mm, and a f-number of 0.68. The full-width half-maximum dimensions at the
geometric focus of this transducer were 0.98 mm, 0.93 mm, and 3.9 mm in the
transverse, elevational, and axial directions, respectively. A custom high-voltage pulser
that generates short therapy pulses of <2 cycles was used to drive the transducer, while
being controlled by a field-programmable gate array (FPGA) board (Altera DE0-Nano
Terasic Technology, Dover, DE, USA) programmed for histotripsy therapy pulsing and
powered by a high voltage DC power supply (GENH750W, TDK-Lambda). During
treatment, the transducer was positioned underneath a custom-designed surgical
mouse stage (Fig.1A) ensuring its submersion in the tank of degassed water. A
68
computer-guided positioning system on a 3-D axis with a 0.05 mm motor resolution was
attached to the transducer in order to run the automated volumetric treatments using a
custom user-interface operated through MATLAB. A linear ultrasound imaging probe
with a frequency range from 10-18 MHz (L18-10L30H-4, Telemed, Lithuania, EU) was
coaxially aligned within the transducer for real-time treatment guidance and monitoring.
This treatment set-up has previously been used for treating subcutaneous and
orthotopic tumors with histotripsy in small animal models [15, 17, 40-43], including our
recent study testing the acute feasibility of ablating CC tumors in the PDX CC tumor
model [37].
3.4.3 Histotripsy Treatment Procedures
Mice were treated when the average cohort tumor diameter was approximately
0.4cm in diameter, to ensure that the smallest tumors were large enough for targeting,
at least 0.3cm in the shortest diameter. Prior to treatment, the water tanks were filled
and degassed, and a water heater was used to maintain the water temperature at
37+4oC. The area of the mouse flank that contained the tumors had fur removed by
applying Nair (Naircare, Ewing, NJ) to the area for 2-4 minutes then washing the area
clean with water and paper towels. Mice were anesthetized with vaporized isoflurane (1-
3% with 1.5 L/min O2 flow rate) and the tumor was submerged in the water through an
opening in the surgical stage. Ultrasound guidance was used to locate the tumor and
confirm the location of the bubble cloud within the tumor throughout the duration of the
treatments (Fig.1C). Using the automated volumetric code to control treatments, an
69
ellipsoid was overlaid onto the tumor, and divided into disks set 0.25mm apart. Points
within each of these disks were set 0.25 mm apart across the entire tumor and 0.5 mm
apart through the depth in the axial direction (Fig.1B). Each point in the disks was
treated with a dose of 500 histotripsy pulses using a pulse repetition frequency (PRF) of
500 Hz and a dwell time of 1 second. The treatment was programmed to move from the
center disk to the external disk, and the same process was then repeated for the
second half of the tumor. For each disk, the treatment started at the central point and
then raster scanned one half and then returned to the center for the second half, to
ensure maximum ablative efficacy.
3.4.4 Pilot Studies to Confirm Ablation Dose and Treatment Margins
Two pilot studies were conducted in order to establish the treatment dose and
margins to be used for chronic experiments. First, negative margin treatments (Fig.2)
were used in an acute pilot study to reconfirm that the dose determined in our prior CC
PDX tumor study was appropriate for this cohort of PDX tumors (n=3) [37]. For these
animals, euthanasia was preformed within 30 minutes after treatment. To establish the
potential of a complete tumor ablation, a second pilot study was conducted testing
positive-margin treatments (Fig.2) which included a ~1-2mm positive margin around the
tumor (n=2) regardless of inclusion of off-target structures. To minimize damage to off
target-tissues (i.e. muscles, skin, and intestines), the therapy transducer was manually
turned off for any bubble cloud that was visually found on ultrasound imaging to be
100% outside of the tumor margins during treatment. Animals in this group were
70
allowed to recover from treatment and were euthanized when it was determined that
pain thresholds were reached with no signs of improvement after subsequent 4-hour
checks, as guided by our IACUC protocol. The symptoms evaluated for these
assessments included lethargy, decreased socialization, difficulty rearing, decreased
extremity strength, squinting, ruffling, as well as irregular respiratory rate and rhythm.
Figure 3-1 Rodent system treatment set up (A). Ultrasound imaging confirmed presence of bubble cloud (indicated by the orange arrow) within CC tumors during treatment (B). For treatment automation, each tumor was defined as a spheroid divided into disks (C). Each treatment disk consisted of a series of points that were used for raster scanning (D). White arrow in (D) represents the direction of therapeutic ultrasound wave propagation.
71
3.4.5 Survival Study
A survival study was conducted to compare histotripsy ablation of CC tumors
between treated and untreated animals (n=6 each). For the survival study, treatment
was applied using a zero-margin method to achieve as close to 100% of the tumor
being targeted with histotripsy as possible while avoiding internal structures such as
intestines being treated around the tumor as was observed in the positive-margin pilot
studies. This zero-margin treatment (Fig.2) led to approximately ~80-90% of the tumor
being targeted. Due tumor shape irregularities, swelling during treatment, and minor
animal movements, some variation was observed in the extent of tumor ablation across
the 6 treated animals. To minimize off target damage, the therapy was manually turned
off for points at which the bubble cloud was visualized to form completely outside of the
tumor. Given the margins used in this cohort, this did not occur often. After treatment,
the animals were recovered and monitored for health responses over the following
weeks. Animals were euthanized when they reached survival endpoints which consisted
of tumors exceeding 1.4 cm in calculated diameter; debilitating health noted including
irregular respiratory rate and rhythm, decreased activity, decreased socialization, or
difficulty moving extremities; or when the IACUC approved end of study time point of 60
days was reached. Progression free survival was defined as growing beyond 20% of the
average tumor diameter that was measured on the day of treatment.
72
Figure 3-2 Schematic showing the planned ablation volumes treatments utilized for this study. Negative-margin treatments were set to have the entire treatment contained within the tumor; zero-margin treatments were set to treat as close to 100% of the tumor as possible without including a significant amount of non-tumor tissue; and positive-margin treatments were set to treat the entire tumor along with sufficient margin that encompassed enough non-tumor tissue to ensure complete ablation of the entire tumor. Arrows indicate tumor tissue not targeted by each planned ablation zone while * indicate intestinal tissue within the treatment margin. Figure created using Biorender.com.
3.4.6 Necropsy and Histopathology
All animals in the positive-margin pilot study and the survival study were
euthanized and necropsied as the animals individually reached the survival end points
described above. During necropsy, all tissues within abdominal cavity as well as nearby
skin and muscles were examined for abnormalities that could have been caused by
treatment, including but not limited to bruising, lacerations, swelling, and deformities.
The tumor, lungs, and liver were collected from all animals and fixed in 10% formalin.
These tissues were then paraffin-embedded and stained with hematoxylin and eosin
(H&E) following standard protocols. Evaluations of tumor tissue to evaluate treatment as
well as the lungs and liver for metastasis were performed by trained individuals and
independently verified by a blinded, board-certified pathologist (D.G.).
73
3.4.7 Statistics
Tumor growth and survival data were analyzed using GraphPad Prism, version 8.
A Student’s two-tailed t-test was used when comparing the survival study experimental
groups tumor growth. Standard Kaplan-Meyer analysis was conducted on progression
free and overall survival. Progression free survival was defined as the tumor exceeding
120% the average diameter on treatment day and overall survival was determined
based on morbidity, as described above. Statistical significance was defined as p ≤ 0.05.
Data are represented as the mean +/- SEM.
Figure 3-3 Histological examination revealed undisturbed tumor stroma in acute untreated mice (A) and completely acellular targeted regions in treated tumors (B). Comparing to before treatment (C), the hypoechoic regions were identifiable in the ultrasound imaging after treatment (D). The treatment area, represented by the dotted yellow circle, closely matched the negative-margin planned ablation volume on ultrasound imaging.
74
3.5 Results
3.5.1 Pilot Studies of Ablation Dose and Margins
Figure 3-4 Using positive, more aggressive margins (n=2) there was more significant skin lesions and subcutaneous bruising that extended well beyond the treatment area across the abdomen (A). Darkened, parrelel lesions that correspond to treatment disks are emphasized with purple arrows. At necropsy, these more aggressive treatments revealed free fluid and intestinal contents within the abdomen and a clear intestinal perforation, indicated by the green arrow (B).
Histotripsy generated a well-confined bubble cloud that destroyed the targeted
CC tumor. Histology results revealed an undisturbed tumor stroma in the untreated mice
(Fig.3A) and a completely acellular region in the treated tumor (Fig.3B). Corresponding
ultrasound guidance for pre-treatment imaging showed the tumor as a hyperechoic
region (Fig.3C), and post-treatment the target region became hypoechoic visualized by
the darker region within the tumor margins (Fig.3D).
When using the positive-margin treatments (n=2), there was no palpable tumor
immediately after treatment, suggesting complete ablation of the entire tumor was
achieved. For both animals treated with the positive-margin ablation volume, significant
side effects were observed including slight bleeding during treatment and post-
treatment skin abrasions that appeared to align with the treatment disks (Fig.4A). In
addition to the skin lesions, there was notable bruising and swelling behind the
75
treatment region, with bruising extending down the flank and across the abdomen.
Immediately after treatment, both animals were found to be bright, alert, and oriented.
At the 12- or 36-hour post-treatment health checks, the mice were able to react to
stimuli normally and had normal range of movement. However, at these checks due to
the increased signs of an adverse event, including lethargic behavior and showed signs
of discomfort and pain, one of the mice were euthanized at each check. Upon necropsy,
it was found that there was free blood and intestinal contents within the abdominal
cavity (Fig.4B). Closer examination found that a loop of small bowel that presided
directly behind the treatment zone within the positive treatment margin had been
perforated in one of the two mice. No other injuries were found. The tumor tissue could
not be harvested for histology from these two animals, given that it appeared to be
completely liquefied and gone at these time points.
3.5.2 Survival Study
Animals that were treated with the zero-margin histotripsy treatments (n=6)
showed no significant skin abrasions during treatment (Fig.5A). During two treatments
small lesions with slight bleeding were noted and completely resolved prior to the
completion of the treatment. Immediately after treatment, there was slight bruising in the
treated areas (Fig.5B). Three days after treatment there was no skin damage in the
untreated mice (Fig.5C) and small, pitted scabs over the treated areas in the histotripsy
treated animals (Fig.5D). Within one week of treatment, these scabs has resolved and
there were no visual differences between the treated and untreated animals. In addition,
76
no signs of pain, distress, nor discomfort were observed after treatment in any of these
animals treated with the zero-margin treatments.
Figure 3-5 No significant skin breaks were observed during treatment (A). Immediately after treatment, there was slight bruising surrounding the tumors, indicated by the blue arrow (B). Three days after treatment there was no skin damage in the untreated mice (C) and small, pitted scabs over the treated areas, noted by the red arrow (D).
Tumor measurements post-treatment were taken with calipers weekly in order to
calculate the average tumor diameter reduction after treatment. Compared to the tumor
diameters in the untreated mice, the diameters of the treated tumors were significantly
reduced and growth retarded for multiple weeks post-treatment (Fig.6A), with a
maximum reduction in tumor size of 73% (p<0.05). Looking at individual mice, it can be
seen that 3 of the 6 of the treated tumors reached a point where the tumor was no
B A
D C
77
longer identifiable approximately 2.5 weeks after treatment (Fig.6B). However, by the
end of the 60-day study, all mice in the treatment group had regrowth of their tumors.
Quantifying the effects of the tumor growth retardation showed that the progression free
and overall survival was significantly higher for the treated mice versus the untreated
(Fig.7). The untreated mice saw tumor growth progression on average by 21 days post
treatment and reached survival end points on average 40 days after treatment. For the
treated mice, progression of tumor growth was prevented until 40 days and survival at
day 60. Of the mice that were treated, 3 of the 6 mice did not reach survival endpoints
by the end of the study.
Figure 3-6 Average tumor diameter (mean + SEM) reduction after treatment, after treatment the difference between groups was significant at every time point (* p<0.05) (A). Individual treated and untreated mouse tumor measurements, emphasizing that multiple treated animals reached a tumor diameter of zero (B). Vertical dotted line emphasizes treatment on day 8. Horizontal dotted line at 14mm emphasizes maximum tumor diameter that any individual animal was allowed to reach in this study based on our IACUC protocols.
At survival time points, the untreated (Fig.8A) and treated (Fig.8B) tumors were
comparable in histologic state. In general, the tumors from all the mice maintained the
features that were consistent with the diagnosed adenosquamous carcinoma from the
78
human donor. At the end of the study, the treated tumors had regrown, and displayed a
comparable level of fibrosis, keratin deposits, and mucin producing areas to the
untreated tumors. One notable difference between the two groups was that the
untreated tumors had larger calcium deposits within the tumors. There was no sign of
metastatic disease in the abdominal and thoracic cavities of any animal, nor did any
animal have signs of splenomegaly. For both the chronic treated and untreated mice the
liver (Fig.8C-D) and lung (Fig.8E-F) parenchyma were benign, with no gross or
histological signs of metastatic disease.
Figure 3-7 Progression free and overall survival increased (p=0.09 and 0.03 respectively) in treatment group compared to untreated. Progression free survival was defined as surpassing 120% of the average tumor diameter the day of treatment (5.29 mm). Vertical dotted line emphasizes treatment on day 8.
79
Figure 3-8 Histological examination revealed undisturbed tumor stroma in chronic (A) untreated mice and tumor regrowth in chronic treated tumors (B). The liver (C-D) and lung (E-F) parenchyma in both untreated (E,C) and treated (D,F) chronic mice were found to be benign. Scale bar = 500um.
3.6 Discussion
In this study, the effects of histotripsy on the treatment of CC in an acute and
survival PDX model was investigated. Results from acute experiments showed that
80
histotripsy generated defined bubble clouds that resulted in a large portion of the CC
tumors being destroyed using a treatment dose of 500 pulses/point (Fig.2), matching
our previous work establishing the feasibility of treating CC tumors with histotripsy [37].
This study was conducted in addition to our previous study that investigated the dose
needed to treat CC PDX tumors in mice with histotripsy in order to confirm this ablation
dose was effective in the new cohort of PDX mice since prior studies of PDX models
have shown significant differences in the tumor microenvironment between cohorts of
PDX mice derived from different patients [39, 44]. It is worth noting that the 500
pulses/point treatment dose used in this study was significantly higher than the 50
pulses/point dose used to achieve a similar level of tumor ablation and growth
retardation in a prior study of hepatocellular carcinoma tumors [17]. As discussed in our
previous study [37], this higher treatment dose for the CC tumors is likely due to the
higher Youngs modulus (56.9±25.6 kPa) [38] for CC tumors compared to hepatocellular
carcinoma tumors (14.86±10.0 kPa) In the prior study of histotripsy for treating
subcutaneous hepatocellular carcinoma tumors, the use of positive margins led to a
report of damage to muscles that were near the tumor [17]. Similarly, the results from
our study demonstrated that histotripsy could not be applied with positive margins
around the tumor without debilitating side effects, resulting in the need for more
conservative margins being used for the survival experiments in this study. More
specifically, results from the positive-margin treatments showed significant damage to
the intestines (Fig.3B) as well as signs of significant pain and distress after treatment. It
is likely that the higher treatment dose of 500 pulses/point needed to ablate CC tumors
increase the extent of damage to the intestines, suggesting that the tissue selective
81
features of histotripsy may be less significant when treating fibrous tumors such as CC,
although further work will be needed to fully investigate this possibility. Overall, the
findings from the initial pilot studies demonstrated the need for more conservative zero-
margin treatments to be used to study the long-term effects of treating CC tumors with
histotripsy. In the animals that were treated with zero-margins, there were no negative
side effects noted beyond mild bruising and scabbing in the days immediately following
treatment (Fig.4).
Results from the survival study showed histotripsy significantly reduced tumor
burden and expanded survival measures compared to control mice. Post-treatment
measurements showed a significant and steady decrease in tumor volume compared to
the untreated groups (Fig.5) for a period of ~3.5 weeks after treatment. The treated
mice survived the histotripsy treatments when utilizing a zero-margin protocol so that
the vast majority of the tumor was treated while avoiding off-target damage, such as
bowel or intestine perforations. Given that these mice were immunocompromised and
therefore lack the immune response necessary to clear any remaining viable cells, it is
not surprising that all treated mice tumor showed tumor regrowth occurred from
surviving cancer cells in the same location as the primary tumor, as expected based on
the zero-margin treatments which did not cover 100% of the tumor volume. This
regrowth also occurred within the 3 mice that reached a point were no tumor mass
palpable after ~3-4 weeks (Fig.5B). This finding was similar to a prior study on using a
comparable histotripsy system to treat hepatocellular carcinoma, where 2/6 of their
treated mice saw tumor size completely deplete before regrowing in
immunocompromised mice [17].
82
Overall, the results of this study continue to support our hypothesis that
histotripsy has the potential to be a viable treatment method for CC, representing the
first completely non-invasive and non-thermal ablation method for this application.
However, results also demonstrate limitations with the animal models used in this study.
For instance, looking to the future, with how slowly these PDX tumors grew (Fig.5), the
lack of metastasis (Fig.7), and the histological features that are similar to human tumors
[45], this model would be useful for studying the possibility of using histotripsy to non-
invasively manage disease through multiple treatments of recurring tumors. Additionally,
in this study we were limited to a safety margin that prevented aggressive margins that
would have the potential to completely eliminate the tumors. When translating this
consideration to larger animal pre-clinical work or human trials, it should be noted that
these off-target damages have been established in the murine model. Future studies
would be needed to determine the dose safety between tumors and margin, critical
structures, such as the intestines. While this study suggests that a complete ablation
may be limited due to safety concerns, overall, these results suggest that histotripsy
would be a usefully palliative option for patients that do not qualify for curative surgeries.
3.7 Conclusion
This study demonstrates the potential to utilize histotripsy in the treatment of CC,
with results showing a maximum average tumor diameter reduction of 73% in a
subcutaneous CC model after 26 days. There were no significant side effects using the
high dose and conservative margins (n=6), with 3 of 6 treated tumors reduced in
83
diameter to an undetectable size in 2.5 weeks of treatment as the ablation homogenate
was reabsorbed. The treated animals were found to have significantly increased
progression-free and overall survival, and no metastases were observed in any animals.
Additionally, due to the subcutaneous placement of the tumors in this study, we were
limited to a safety margin that prevented aggressive, positive margins that would have
the potential to completely eliminate the tumors in all animals. Overall, the results of this
study support the hypothesis that histotripsy has the potential to be a viable treatment
method for CC, representing the first completely non-invasive and non-thermal ablation
method for this application.
3.8 Acknowledgment
This research was supported by a grant from the Focused Ultrasound
Foundation (FUSF-RAP-S-18-00011) and was conducted in memory of Andrew J.
Lockhart, who passed away from cholangiocarcinoma on September 30, 2016. The
authors encourage anyone interested to consider a donation to the Focused Ultrasound
Foundation in Andrew’s name.
The authors would also like to thank the ICTAS Center for Engineered Health, the
Virginia Tech Department of Biomedical Engineering and Mechanics, Virginia Tech
Carilion School of Medicine, and the Virginia-Maryland College of Veterinary Medicine
for their support of this study.
84
3.9 References
[1] ACS, "Cancer Facts & Figures 2020," American Cancer Society, 2020.
[2] E. U. Cidon, "Resectable cholangiocarcinoma: reviewing the role of adjuvant
strategies," Clinical Medicine Insights: Oncology, vol. 10, p. CMO. S32821, 2016.
[3] M. Jansen, R. Van Hillegersberg, R. Chamuleau, O. Van Delden, D. Gouma, and
T. Van Gulik, "Outcome of regional and local ablative therapies for hepatocellular
carcinoma: a collective review," European Journal of Surgical Oncology (EJSO),
vol. 31, no. 4, pp. 331-347, 2005.
[4] L. Rossi et al., "Current approach in the treatment of hepatocellular carcinoma,"
World journal of gastrointestinal oncology, vol. 2, no. 9, p. 348, 2010.
[5] S. K. Venkatesh et al., "MR elastography of liver tumors: preliminary results,"
American Journal of Roentgenology, vol. 190, no. 6, pp. 1534-1540, 2008.
[6] D. M. Nagorney, J. H. Donohue, M. B. Farnell, C. D. Schleck, and D. M. Ilstrup,
"Outcomes after curative resections of cholangiocarcinoma," Archives of surgery,
vol. 128, no. 8, pp. 871-878, 1993.
[7] K. Feng et al., "A randomized controlled trial of radiofrequency ablation and
surgical resection in the treatment of small hepatocellular carcinoma," Journal of
hepatology, vol. 57, no. 4, pp. 794-802, Oct 2012, doi:
10.1016/j.jhep.2012.05.007.
[8] M. S. Chen et al., "A prospective randomized trial comparing percutaneous local
ablative therapy and partial hepatectomy for small hepatocellular carcinoma," (in
85
eng), Ann Surg, Randomized Controlled Trial, Research Support, Non-U.S. Gov't
vol. 243, no. 3, pp. 321-8, Mar 2006, doi: 10.1097/01.sla.0000201480.65519.b8.
[9] R. Salem et al., "Institutional Decision to Adopt Y90 as Primary Treatment for
HCC Informed by a 1,000-patient 15-year Experience," Hepatology, Dec 1 2017,
doi: 10.1002/hep.29691.
[10] A. Forner, J. M. Llovet, and J. Bruix, "Hepatocellular carcinoma," Lancet, vol.
379, no. 9822, pp. 1245-55, Mar 31 2012, doi: 10.1016/S0140-6736(11)61347-0.
[11] L. European Association for the Study of the, R. European Organisation for, and
C. Treatment of, "EASL–EORTC Clinical Practice Guidelines: Management of
hepatocellular carcinoma," Journal of hepatology, vol. 56, no. 4, pp. 908-943,
2012. [Online]. Available:
http://linkinghub.elsevier.com/retrieve/pii/S0168827811008737?showall=true.
[12] A. Gillams et al., "Thermal ablation of colorectal liver metastases: a position
paper by an international panel of ablation experts, The Interventional Oncology
Sans Frontieres meeting 2013," Eur Radiol, vol. 25, no. 12, pp. 3438-54, Dec
2015, doi: 10.1007/s00330-015-3779-z.
[13] K. Han, H. K. Ko, K. W. Kim, H. J. Won, Y. M. Shin, and P. N. Kim,
"Radiofrequency ablation in the treatment of unresectable intrahepatic
cholangiocarcinoma: systematic review and meta-analysis," Journal of Vascular
and Interventional Radiology, vol. 26, no. 7, pp. 943-948, 2015.
[14] E. A. Takahashi et al., "Thermal ablation of intrahepatic cholangiocarcinoma:
safety, efficacy, and factors affecting local tumor progression," Abdominal
Radiology, vol. 43, no. 12, pp. 3487-3492, 2018.
86
[15] E. Vlaisavljevich et al., "Non-Invasive Ultrasound Liver Ablation Using Histotripsy:
Chronic Study in an In Vivo Rodent Model," Ultrasound Med Biol, vol. 42, no. 8,
pp. 1890-902, Aug 2016, doi: 10.1016/j.ultrasmedbio.2016.03.018.
[16] E. Vlaisavljevich et al., "Image-guided non-invasive ultrasound liver ablation
using histotripsy: Feasibility study in an in vivo porcine model," Ultrasound in
medicine & biology, vol. 39, no. 8, pp. 1398-1409, 2013.
[17] T. Worlikar et al., "Histotripsy for Non-Invasive Ablation of Hepatocellular
Carcinoma (HCC) Tumor in a Subcutaneous Xenograft Murine Model," in 2018
40th Annual International Conference of the IEEE Engineering in Medicine and
Biology Society (EMBC), 2018: IEEE, pp. 6064-6067.
[18] A. D. Maxwell, C. A. Cain, T. L. Hall, J. B. Fowlkes, and Z. Xu, "Probability of
cavitation for single ultrasound pulses applied to tissues and tissue-mimicking
materials," Ultrasound in medicine & biology, vol. 39, no. 3, pp. 449-465, 2013.
[19] E. Vlaisavljevich et al., "Effects of Ultrasound Frequency and Tissue Stiffness on
the Histotripsy Intrinsic Threshold for Cavitation," Ultrasound Med Biol, 2015.
[20] Z. Xu et al., "Controlled ultrasound tissue erosion," IEEE transactions on
ultrasonics, ferroelectrics, and frequency control, vol. 51, no. 6, pp. 726-736,
2004.
[21] Z. Xu, M. Raghavan, T. L. Hall, M.-A. Mycek, J. B. Fowlkes, and C. A. Cain,
"Evolution of bubble clouds produced in pulsed cavitational ultrasound therapy -
histotripsy," IEEE Trans Ultrason Ferroelectr Freq Control., vol. 55, no. 5, pp.
1122-1132, 2008.
87
[22] E. Vlaisavljevich, A. Maxwell, M. Warnez, E. Johnsen, C. Cain, and Z. Xu,
"Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
mechanical properties," (in eng), IEEE Transactions on Ultrasonics,
Ferroelectrics, and Frequency Control, vol. 61, no. 2, pp. 341-52, Feb 2014, doi:
10.1109/TUFFC.2014.6722618.
[23] E. Vlaisavljevich, A. Maxwell, L. Mancia, E. Johnsen, C. Cain, and Z. Xu,
"Visualizing the Histotripsy Process: Bubble Cloud-Cancer Cell Interactions in a
Tissue-Mimicking Environment," (in Eng), Ultrasound Med Biol, Jul 8 2016, doi:
S0301-5629(16)30099-0 [pii] 10.1016/j.ultrasmedbio.2016.05.018.
[24] K. C. Longo et al., "Histotripsy Ablations in a Porcine Liver Model: Feasibility of
Respiratory Motion Compensation by Alteration of the Ablation Zone Prescription
Shape," CardioVascular and Interventional Radiology, pp. 1-7, 2020.
[25] T. Worlikar et al., "Non-Invasive Orthotopic Liver Tumor Ablation Using
Histotripsy in an in Vivo Rodent Hepatocellular Carcinoma (HCC) Model,"
presented at the Society of Interventional Oncology, Boston, MA, USA, 2019.
[26] K. C. Longo et al., "Robotically Assisted Sonic Therapy (RAST) for Noninvasive
Hepatic Ablation in a Porcine Model: Mitigation of Body Wall Damage with a
Modified Pulse Sequence," Cardiovasc Intervent Radiol, vol. 42, no. 7, pp. 1016-
1023, Jul 2019, doi: 10.1007/s00270-019-02215-8.
[27] E. Vlaisavljevich et al., "Non-invasive liver ablation using histotripsy: preclinical
safety study in an in vivo porcine model," Ultrasound in medicine & biology, vol.
43, no. 6, pp. 1237-1251, 2017.
88
[28] T. Worlikar et al., "Non-invasive liver cancer ablation using histotripsy in an in
vivo murine hepatocellular carcinoma (HCC) model," in Ultrasonics Symposium
(IUS), 2017 IEEE International, 2017: IEEE, pp. 1-1.
[29] J. Vidal Jove et al., "Phase I Study of Safety and Efficacy of Hepatic Histotripsy:
Preliminary Results of First in Man Experience with Robotically-Assisted Sonic
Therapy," presented at the International Society of Therapeutic Ultrasound,
Barcelona, Spain, 2019.
[30] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," Physics in Medicine & Biology, vol. 59, no. 2, p. 253, 2013.
[31] E. Vlaisavljevich, Z. Xu, A. Arvidson, L. Jin, W. Roberts, and C. Cain, "Effects of
thermal preconditioning on tissue susceptibility to histotripsy," Ultrasound in
medicine & biology, vol. 41, no. 11, pp. 2938-2954, 2015.
[32] T. L. Hall, C. R. Hempel, K. Wojno, Z. Xu, C. A. Cain, and W. W. Roberts,
"Histotripsy of the prostate: dose effects in a chronic canine model," Urology, vol.
74, no. 4, pp. 932-937, 2009.
[33] A. Lake, Z. Xu, J. Wilkinson, C. Cain, and W. Roberts, "Renal ablation by
histotripsy—does it spare the collecting system?," The Journal of urology, vol.
179, no. 3, pp. 1150-1154, 2008.
[34] C. R. Hempel, T. L. Hall, C. A. Cain, J. B. Fowlkes, Z. Xu, and W. W. Roberts,
"Histotripsy fractionation of prostate tissue: local effects and systemic response
in a canine model," The Journal of urology, vol. 185, no. 4, pp. 1484-1489, 2011.
89
[35] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," (in eng), Phys Med Biol, vol. 59, no. 2, pp. 253-70, Jan 20 2014, doi:
10.1088/0031-9155/59/2/253.
[36] V. A. Khokhlova et al., "Histotripsy methods in mechanical disintegration of
tissue: Towards clinical applications," International journal of hyperthermia, vol.
31, no. 2, pp. 145-162, 2015.
[37] A. Hendricks et al., "Histotripsy for the Treatment of Cholangiocarcinoma Liver
Tumors: In Vivo Feasibility and Ex Vivo Dosimetry Study," IEEE Transactions on
Ultrasonics, Ferroelectrics, and Frequency Control, 2021.
[38] A. Guibal et al., "Evaluation of shearwave elastography for the characterisation of
focal liver lesions on ultrasound," European radiology, vol. 23, no. 4, pp. 1138-
1149, 2013.
[39] R. M. Brock et al., "Patient derived xenografts expand human primary pancreatic
tumor tissue availability for ex vivo irreversible electroporation testing," Frontiers
in Oncology, vol. 10, 2020.
[40] A. Hendricks et al., "Histotripsy initiates local and systemic immunological
response and reduces tumor burden in breast cancer
AD Hendricks, J Howell, R Schmieley, S Kozlov, A Simon, ..." The Journal of
Immunology, vol. 202, 2019.
[41] A. D. Hendricks et al., "Determining the mechanism of the immune response to
histotripsy ablation of pancreatic cancer," ed: Am Assoc Immnol, 2020.
90
[42] S. Qu et al., "Non-thermal histotripsy tumor ablation promotes abscopal immune
responses that enhance cancer immunotherapy," Journal for ImmunoTherapy of
Cancer, vol. 8, no. 1, 2020.
[43] A. Hendricks-Wenger et al., "Histotripsy Ablation Alters the Tumor
Microenvironment and Promotes Immune System Activation in a Subcutaneous
Model of Pancreatic Cancer," IEEE Transactions on Ultrasonics, Ferroelectrics,
and Frequency Control, 2021.
[44] L. E. Dobrolecki et al., "Patient-derived xenograft (PDX) models in basic and
translational breast cancer research," Cancer and Metastasis Reviews, vol. 35,
no. 4, pp. 547-573, 2016.
[45] B. Blechacz, M. Komuta, T. Roskams, and G. J. Gores, "Clinical diagnosis and
staging of cholangiocarcinoma," Nature reviews Gastroenterology & hepatology,
vol. 8, no. 9, pp. 512-522, 2011.
91
Chapter 4. Histotripsy Ablation in Preclinical Animal Models of Cancer and Spontaneous Tumors in Veterinary Patients
Alissa Hendricks-Wenger, Lauren Arnold, Jessica Gannon, Alex Simon, Neha Singh, Hannah Sheppard, Margaret A. Nagai-Singer, Khan Mohammad Imran1, Kiho Lee,
Sherrie Clark-Deener, Christopher Byron, Martha M. Larson, John H. Rossmeisl, Sheryl L. Coutermarsh-Ott, Nikolaos Dervisis, Shawna Klahn, Joanne Tuohy, Irving C. Allen,
Eli Vlaisavljevich This chapter is excerpted from a manuscript submitted to IEEE Transactions in UFFC.
92
4.1 Abstract
New therapeutic strategies are direly needed in the fight against cancer. Over the last
decade, several focal tumor ablation strategies have emerged as stand-alone or
combination therapies. Histotripsy, an ultrasound ablation method that destroys tissue
through the precise control of acoustic cavitation, has emerged as the first completely
non-invasive, non-thermal, and non-ionizing ablation method for the treatment of
cancer. Histotripsy does not share the limitations of thermal ablation, can produce
consistent and rapid ablations, and even near critical structures. Additional benefits
include real-time image-guidance, high precision, and the ability to treat tumors of any
predetermined size and shape. Unfortunately, the lack of clinically and physiologically
relevant pre-clinical cancer models is often a significant limitation with all focal tumor
ablation strategies. The majority of studies testing histotripsy for cancer treatment have
focused on small animal models, which have been critical in moving this field forward
and will continue to be essential for providing mechanistic insight. However, these
models have significant limitations in terms of scale and translational value. To address
these limitations, a diverse range of large animal models and spontaneous tumor
studies in veterinary patients have emerged to complement existing rodent models.
These models and veterinary patients have proven to be excellent at providing realistic
models for developing and testing histotripsy devices and techniques designed for
future use in human patients. Here, we compare histotripsy ablation across a
representative spectrum of preclinical animal models and spontaneous tumors in
93
veterinary patients. Together, these studies demonstrate the utility and advantages of
these bench-to-kennel-to-bedside approaches in the development of histotripsy.
94
4.2 Introduction
Histotripsy is a focused ultrasound ablation method that destroys tissue through the
precise control of acoustic cavitation [1-4]. Using microsecond long, high-pressure
pulses applied by a focused ultrasound transducer coupled to the patient, a histotripsy
bubble cloud is generated non-invasively inside the targeted tissue [5-9]. The rapid
expansion and collapse of these bubbles result in complete disruption of the target
tissue into acellular debris [10]. As a non-thermal ablation method, histotripsy has
shown the potential to overcome the limitations associated with thermal ablation. For
instance, histotripsy has been shown to produce consistent and complete ablation in the
liver, which is highly vascular, when applied through abdominal or transcostal acoustic
windows [1, 3, 11, 12]. Histotripsy has also shown the ability to ablate tissue near vital
structures such as major vessels, bile ducts, and nerves while preserving these
structures [1, 3, 9, 13, 14]. Due to these features, histotripsy is currently being
developed for multiple clinical applications, most notably for the treatment of cancer [4,
15-17]. However, despite the promise of histotripsy and the many positive results
reported in preclinical studies, to date, only a single human clinical trial has been
conducted to test histotripsy for the treatment of cancer [18]. This recent Phase I clinical
trial was conducted in non-curative patients with multifocal primary and metastatic liver
tumors, with results showing histotripsy could safely target and ablate targeted liver
tumors [18]. To our knowledge, no other human clinical trials of histotripsy for the
95
treatment of cancer have been conducted, limiting the potential widespread translation
of histotripsy into clinical practice as an improved non-invasive tumor ablation method.
Similar to other focal tumor ablation modalities, one of the primary factors limiting the
rapid translation of histotripsy into the clinic has been the lack of appropriate animal
models for studying the safety, effectiveness, and optimization of histotripsy for the
treatment of different cancers. Previous efforts to develop histotripsy for specific
oncology applications have been split into separate studies testing human-scale
histotripsy systems for targeting and ablating healthy tissue in large animal models that
have similar anatomy and physiology to human patients (primarily pigs) [1, 3, 11, 12].
These studies are typically complemented by separately testing histotripsy tumor
ablation in rodents (mice, rats) using customized small animal devices that allow
histotripsy to be applied in this setting, albeit not using the systems or acoustic
parameters that will ultimately be developed for use in humans [2, 4, 19, 20].
Murine models are the dominant in vivo model in the biomedical engineering and
cancer biology fields and are essential in acquiring fundamental data. Mice are easy to
house and handle, relatively inexpensive, share many genetic similarities to humans,
and there are many transgenic and knock-out lines available to study specific disease
mechanisms, proteins, and pathways [21-26]. In typical mouse models used to study
tumor ablation modalities, the vast majority of studies are mouse or human cell line
based. While these models offer excellent reproducibility and standardization, they
rarely recapitulate the clinical, in situ, microenvironment found in patients or critical
biological hallmarks of the original tumor of origin. This is especially true in
subcutaneous, flank generated tumors. Orthotopic tumors that are transplanted into the
96
organ of interest can provide improved insight but still suffer from cell line specific
limitations related to cancer biology. Spontaneous and induced tumors in small animals
are more physiologically relevant to human patients. However, the inability to predict
tumor development, location, progression, and lack of reproducibility are all limitations
to these models. Likewise, the lack of translational acoustic windows, limited imaging,
lack of precision, and lack of physiological relevance are all major limitations of small
animal models in the development of histotripsy and other focused ultrasound
modalities. While critical for proof-of-concept testing and mechanistic insight, the
miniaturization of the model has significantly slowed the rate of clinical translation and
made it difficult to reliably test the effectiveness of clinical prototype histotripsy systems
for imaging, targeting, and effectively treating different tumors types. As a result, there is
a significant need for improved preclinical large animal tumor models for studying
histotripsy tumor ablation using clinically relevant devices and treatment parameters to
enable the more rapid and successful translation of histotripsy into clinical practice for
the treatment of cancer in humans.
Large animal models can address many of the shortcomings often cited for
rodents, in many cases offering direct translation from animal to human. For example,
the same tumor ablation systems can typically be utilized, similar clinical/surgical
techniques can be refined, and many large animal models accurately recapitulate
human anatomical and physiological features. This is critical for imaging and
ultrasound-based modalities such as histotripsy. However, the increased size and
complexity of the species results in several significant limitations typically associated
with cost, the requirement for specialized housing, and the need for unique institutional
97
resources. Most large animal models are based on spontaneous tumor development,
which has some of the same limitations commonly cited for small animal studies.
Recent technological advancements in gene editing technologies have largely negated
these issues, specifically in pig models. In general, pigs have yet to play a significant
role in experimental oncology [27]. However, with the rise of technologies capable of
inducing genetic modifications in large mammals, such as CRISPRs and TALONs, there
is a renewed interest in the pig as a cancer model. This is especially true in the area of
medical devices, where the utilization of the same devices, instruments, and clinical
practice can be utilized in pigs and humans, enabling a more rapid translation to clinical
trials. Over the last decade, genetically modified porcine cancer models have been
generated targeting APC, TP53, KRAS, GLI2, V-H-RAS, and BRACA1[27]. Indeed, the
recently developed “Oncopig” cancer model (OCM) has the potential to bring the pig to
the forefront of tumor models [28]. These novel transgenic swine models recapitulate
human cancer through Cre recombinase induced site and cell-type specific expression
of KRAS and TP53 transgenes [29]. To express Cre, pigs must be either crossed with
transgenic animals expressing cell type specific Cre or direct injection into the tissue
type of interest with an adenoviral vector encoding Cre recombinase [28]. Both KRAS
and TP53 mutations are common in multiple cancers and multiple OCM models appear
to be currently under development [28].
The OCM model is expected to have several strengths, including an accurate
recapitulation of the tumor microenvironment, accurate immune system profile, and the
outbred animals are expected to better model human patients [28]. However, the
reliance on Cre recombinase and associated delivery vectors will result in an
98
unpredictable number of tumors that form in random locations. Likewise, while tumor
development in the OCM model is rapid, it is highly difficult to predict tumor progression
in this model. This can add significant cost and time. As an alternative strategy,
research teams have focused on immunocompromised pig models, where deletions in
the RAG2 and IL2RG genes have generated animals with similar characteristics to NOD
scid gamma (NSG) mice, which are susceptible to human tissue engraftment [30].
Similar to NSG mice, human tumor cell lines, and patient-derived xenograft (PDX)
tissue can be engrafted. This allows a highly predictable tumor that can be surgically
placed subcutaneously or orthotopically in a human-sized animal with similar anatomy
and physiology. Tumors can be expanded to increase tissue availability for ex vivo
studies or surgically placed near critical structural elements (such as blood vessels or
nerve bundles) to evaluate and model tumor ablation in hard to treat locations within the
organ. Another advantage of this strategy is the use of human cells in a pig model. This
allows robust identification, evaluation, and quantification of the ablation zone and
metastatic tumors at the histopathologic level using human antibodies in porcine
tissues. The lack of a functional immune system is a clear disadvantage of
immunocompromised pig models as they require housing under highly specialized
germfree conditions that are only available at a limited number of institutions.
Beyond tumor models, spontaneous cancers in pet animals offer unique
translational research opportunities for cancer imaging and device development that
provide a robust combination of clinical and physiological relevance to humans. Indeed,
comparative oncology studies have revealed significant similarities between many
canine and human cancers [31, 32]. Companion animals, like humans, are genetically
99
diverse and are exposed to many of the same environmental influences as their owners.
Canine cancer patients have been successfully used in numerous studies to provide
translatable pharmacokinetic and pharmacodynamic data [33-38], and these trials have
been critical for determining the scheduling/dosing of small molecular inhibitors in
humans [39]. The spontaneous nature of canine cancers, their biologic similarities to
human cancers, and the anatomic similarities between dogs and people offer highly
relevant clinical conditions for device development.
In this work, we provide an overview of small animal tumor models currently being
used for histotripsy studies, as well as, emerging large animal models under
development by our group for studying histotripsy tumor ablation. In the first part of this
work, we provide a comparison of the histotripsy experimental set-ups commonly used
in small and large animal experiments. We then show representative results of
histotripsy tumor ablation in small animal (mice) cancer models including examples of
subcutaneous, orthotopic, and patient-derived xenograft (PDX) models. Next, we show
results from a recently developed immunocompromised RAG2/IL2RG deficient pig
tumor model that better mimic human patient anatomy and physiology compared to the
mouse models. Finally, we discuss the opportunity to investigate the safety, efficacy,
and optimization of histotripsy in veterinary populations of dogs with spontaneous
tumors and provide an overview of the canine oncology patients at our location as well
as examples of our preliminary studies of histotripsy for the treatment of a variety of
cancers in these veterinary patients. Overall, the results of this study will provide an
overview of the benefits and limitations of currently available animal models and
highlight the potential to utilize emerging large animal tumor models and veterinary
100
cancer patients as a means to enable more rapid and reliable clinical translation of
histotripsy and other emerging FUS methods for the treatment of patients with a wide
range of cancers.
4.3 Procedures
4.3.1 Histotripsy Systems and Pressure Calibrations
Two separate histotripsy systems were used in this study for the small animal and
large animal in vivo histotripsy studies. The small animal in vivo experiments were
conducted using a custom 8-element 1MHz histotripsy transducer with a geometric
focus of 36 mm, an aperture size of 52.7 mm, and an f-number of 0.68 (Fig.1B). The
transducer was driven via a custom high-voltage pulser designed to generate short
therapy pulses of <2 cycles controlled by a field-programmable gate array (FPGA)
board (Altera DE0-Nano Terasic Technology, Dover, DE, USA) programmed for
histotripsy therapy pulsing. The transducer was positioned in a tank of degassed water
beneath a custom-designed mouse surgical stage and attached to a computer-guided
3-D positioning system with 0.05 mm motor resolution to control the automated
volumetric treatments (Fig.1A). A linear ultrasound imaging probe with a frequency
range of 10-18 MHz (L18-10L30H-4, Telemed, Lithuania, EU) was coaxially aligned
inside the transducer for treatment guidance and monitoring (Fig.1C). The transducer
was powered by a high voltage DC power supply (GENH750W, TDK-Lambda), and the
system was controlled using a custom user-interface operated through MATLAB
101
(MathWorks). This transducer design has been used previously for multiple small
animal histotripsy studies in mouse and rat models [2, 4, 19, 20].
Figure 4-1 Example Small Animal Histotripsy System. (A) A small animal system consisting of a 1) 1 MHz therapy transducer with an 2) imaging transducer coaxially aligned and mounted to a 3) 3D motorized positioning system to deliver hisotripsy to target submerged in degassed water on a 5) subject stage. A custom-built electronic driving system is controlled with computer software. (B) Linear ultrasound imaging probe coaxially aligned with 1 MHz therapy transducer used to (C) treat tumors in-vivo. (D) Acoustic waveform for 1 MHz transducer collected with a hydrophone at a peak negative pressure of ~10 MPa. (E) 1 MHz bubble cloud captured on a high-speed camera.
For the large animal in vivo experiments, histotripsy was applied using a custom 32-
element 500 kHz array transducer with a geometric focus of 78 mm, an elevational
aperture size of 112 mm, and a transverse aperture size of 128 mm, with corresponding
f-numbers of 0.61 (transverse) and 0.70 (elevational) (Fig.2B). The transducer was
driven via custom high-voltage pulser designed to generate short therapy pulses of <2
cycles controlled by a field-programmable gate array (FPGA) board (Altera DE0-Nano
Terasic Technology, Dover, DE, USA) programmed for histotripsy therapy pulsing. The
transducer was fixed onto a prototype clinical histotripsy treatment cart (HistoSonics,
Ann Arbor, MI, USA) that included an ultrasound imaging system, robotic micro-
positioner, and customized for applying volumetric histotripsy ablations (Fig.2A). A 3
102
MHz curvilinear imaging probe (Model C5-2, Analogic Corp., Peabody, MA) was
coaxially aligned through a central hole in the therapy transducer in order to allow real-
time treatment guidance and monitoring (Fig.2B). The transducer was powered by a
high voltage DC power supply (GENH750W, TDK-Lambda) and the transducer was
controlled using a custom user-interface operated through MATLAB (MathWorks) which
was triggered from the HistoSonics system. This overall system is similar to the one
used in recent large animal histotripsy studies conducted on healthy pigs as well as a
recent Phase I clinical trial of histotripsy for treating liver cancer patients [1, 18], with the
only difference being the custom 32-element transducer that was used in this study in
place of the clinical histotripsy transducer used in the prior work.
Figure 4-2 Example Large Animal Histotripsy System. (A) Histotripsy system designed for liver cancer ablation consisting of 1) therapy and 2) imaging transducers attached to an 3) articulating arm with a 4) motorized micro-positioner controlled by 5) custom software. (B) Image of therapy transducer with co-axially aligned imaging probe used for (C) previous in vivo liver studies. (D) Acoustic waveform for 500 kHz transducer collected with a hydrophone at a peak negative pressure of ~10 MPa. (E) 500 kHz bubble captured on a high-speed camera.
103
4.3.2 Hydrophone Focal Pressure Measurements, Beam Profiles, and Optical Imaging
Focal pressure waveforms for the 1MHz and 500kHz transducers were measured by
a high sensitivity rod hydrophone (HNR-0500, Onda Corporation, Sunnyvale, CA, USA)
and a custom-built fiber optic probe hydrophone (FOPH) [40, 41] in degassed water at
the focal point of each transducer. The rod hydrophone was used to measure the 1D
focal beam profiles of the transducer in the lateral, elevational, and axial directions at a
peak negative pressure (p-) of ~1.8 MPa. Focal pressures were directly measured with
the FOPH up to a p- of ~20 MPa. Pressure waveforms at higher p- were estimated by a
linear extrapolation [42] since p- values above ~20MPa couldn’t be directly measured
due to cavitation on the tip of the FOPH fiber. All waveforms were measured using a
Tektronix TBS2000 series oscilloscope at a sample rate of 500MS/s, with the
waveforms averaged over 128 pulses and recorded in MATLAB.
The bubble clouds generated by each transducer were characterized using high-
speed optical imaging as described in previous studies [43, 44]. Briefly, the focal region
inside 1% agarose tissue phantoms was imaged after acoustic propagation for each
pulse using the 1 MHz and 500 kHz therapy transducers described above. Optical
imaging was performed using a high-speed camera (FLIR Blackfly S monochrome,
BFS-U3-32S4M-C 3.2 MP, 118 FPS, Sony IMX252, Mono, FLIR Integrated Imaging
Solutions, Richmond, BC, Canada) and a 100 mm F2.8 Macro lens (Tokina AT-X Pro,
Kenko Tokina Co., LTD, Tokyo, Japan). The camera was externally triggered to record
one image per applied pulse, and the tissue phantom was backlit by a custom-built
pulsed white-light LED strobe light capable of high-speed triggering with 1 μs
104
exposures. Strobe duration was kept low (3-5 µs) and all exposures were centered at a
delay of 8.5 µs after pulse arrival at the focus. This timing schema was previously
demonstrated as the optimum delay for visualizing complete bubble clouds [43, 44]. The
bubble cloud dimensions observed on optical imaging were subsequently used to
determine the spacing between adjacent treatment points in in vivo tumor ablation
experiments.
4.3.3 Murine Tumor Models
The ability of histotripsy to ablate tumors in in vivo mouse models was tested using
the 1 MHz small animal histotripsy system to target tumors in three murine cancer
models including 1) a subcutaneous renal cell carcinoma (RCC) model, 2) a patient-
derived xenograft (PDX) subcutaneous cholangiocarcimona model, and 3) a 4T1
orthotopic mammary tumor model. All procedures in this study were conducted in
accordance with the ARRIVE guidelines and approved by the IACUC at Virginia Tech.
In the subcutaneous RCC model, male BALB/cCr mice (n=4) were injected in the right
flank with 6x106 Renca RCC cells (ATCC CRL-2947) in Matrigel. In the subcutaneous
cholangiocarcinoma (CC) PDX model, female Nod scid gamma (NSG) mice (n=4) with
subcutaneous flank PDX CC tumors were commercially acquired from the Jackson
Laboratory (TM01225), through a process previously described [45]. In the orthotopic
mammary tumor model, female BALB/C mice (n=4) were injected with 1.2 X 106 murine
4T1 mammary tumor cells (ATCC CRL-2539) in the abdominal mammary glands. Once
tumors were palpable, animals were monitored three times weekly and tumor diameter
105
was determined by taking two perpendicular measurements, one of which was the
widest point of the tumor. A single tumor diameter was calculated as the square root of
the product of the two measurements. The mice underwent histotripsy treatment when
their tumors reached the targeted treatment size ~0.5-1.0 cm in diameter.
For all tumors, histotripsy was applied after each tumor reached the
predetermined target size. For all treatments, mice were anesthetized with inhaled
isoflurane and secured to a platform over a tank of degassed water, with the
temperature maintained between 35-39°C. Before treatment, fur was removed over the
tumor and the surrounding area by applying Nair (Naircare, Ewing, NJ, USA) for 1-2
minutes and removed by wiping off with a wet paper towel. For treatment, mice were
anesthetized with isoflurane (1.5 L/min oxygen flow rate with 1.0-2.5% isoflurane) and
placed prone on the subject stage with their tumor submerged underwater (Fig.1A).
Animals’ respiratory rate and rhythm were monitored during treatment by trained
personnel. Histotripsy was then applied to a predetermined volume of the tumor using
the custom 1MHz small animal transducer described above and shown in Figure 1.
During treatment, histotripsy was applied using single cycle pulses applied at a pulse
repetition frequency (PRF) of 250 Hz with a 1 second dwell time at each treatment
point. To apply a volumetric treatment, the focus was raster-scanned through a series of
treatment points spaced by 1.5 mm in the axial direction and 0.5 in the lateral for the
Renca and 4T1 tumors. Given that the CC tumors are stiffer and more resistant to
histotripsy, a PRF of 500 Hz with a 1 second dwell time was used to double the per
point dose and the spacing was reduced to 0.75 mm and 0.25 mm to increase the
overlap between adjacent treatment points.
106
4.3.4 RAG2/IL2RG Deficient Porcine Tumor Model
An ideal animal model for studying histotripsy tumor ablation using clinical prototype
systems would 1) be implantable so that specific locations could be chosen for the
hypothesis being tested, 2) allow implantation of human tumor cell lines to closely mimic
the natural history and composition of human tumors, and 3) be in an animal large
enough to evaluate human-scale devices. To accomplish these goals, our team has
recently developed RAG2/IL2RG double knockout pigs [46-52]. Oocytes were aspirated
from sow ovaries and matured in maturation medium (TCM-199 medium supplemented
with 0.5 IU ml−1 FSH, 0.5 IU ml−1 LH, 0.82 mM cysteine, 3.02 mM glucose, 0.91 mM
sodium pyruvate and 10 ng ml−1 EGF) for 40 – 44 hours. After in vitro maturation,
cumulus cells were removed by vortexing in the presence of hyaluronidase, then mature
oocytes extruding the first polar body were transferred to fertilization media (modified
Tris-buffered medium supplemented with 113.1 mM NaCl, 3 mM KCl, 7.5 mM CaCl2,
11 mM glucose, 20 mM Tris, 2 mM caffeine, 5 mM sodium pyruvate and 2 mg ml−1
BSA). Extended semen was washed with PBS, then introduced into the IVF dishes
containing oocytes. The gametes were co-incubated for 5 hours at 38.5 °C and 5%
CO2. Microinjection was conducted in manipulation media (TCM199 with 0.6 mM
NaHCO3, 2.9 mM HEPES, 30 mM NaCl, 10 ng ml−1 gentamicin and 3 mg ml−1 BSA)
covered with mineral oil on the heated stage of a Nikon inverted microscope (Nikon
Corporation, Tokyo, Japan). Two sgRNAs targeting the RAG2 gene, three sgRNAs
targeting the IL2RG gene, and Cas9 mRNA (10 ng/µl sgRNA each and 20 ng/μl Cas9
mRNA) were introduced into the cytoplasm of the presumable zygotes using a FemtoJet
107
microinjector (Eppendorf, Hamburg, Germany). The injected embryos were moved to
culture media supplemented with 10 ng ml−1 GM-CSF and incubated at 38.5 °C, 5%
CO2, and 5% O2 until embryo transfer [53]. The cultured embryos were transferred into
a surrogate gilt at day 4 post-IVF by surgically transferring into the oviduct. Pregnancy
was determined by ultrasound at around day 30 of gestation. To genotype the mutations
generated by CRISPR/Cas9 system, genomic DNA was isolated from tail-tips of the
newborn piglets using PureLink Genomic DNA kit (Thermo Fisher Scientific) following
the manufacturer’s instructions. Primers flanking projected double-strand break (DSB)
sites were designed to amplify the target region of RAG2 and IL2RG genes. The target
regions were PCR amplified using Dream Taq DNA Polymerase (Thermo Fisher
Scientific). PCR conditions were as follows: initial denature at 95 °C for 2 min; denature
at 95 °C for 30 sec, annealing at 60 °C for 30 sec, and extension at 72 °C for 30 sec for
34 cycles; 72 °C for 5 min; and holding at 4 °C. The PCR amplicons were sequenced to
determine the mutation types generated by the CRISPR/Cas9 system.
Immunocompromised state was verified with flow cytometry to analyze the presence of
B (CD79A), T (CD3E), and NK (CD3E-/CD56+) cells.
Pigs were delivered through a sterile hysterectomy process previously described
by a board-certified large animal veterinarian [54]. Pigs were monitored regularly until
fully recovered from anesthesia and able to eat independently. Tail snips were taken
from each piglet while still under the influence of anesthesia for genotyping, and ears
were clipped for numbering. The pigs were born into sterile, gnotobiotic isolators and
were fed sterile boxed milk throughout the study to prevent infections [54].
108
Human Panc01 cells (National Cancer Institute DTP, DCTP Tumor Repository)
were propagated in DMEM supplemented with 10% FBS and 1% penicillin/
streptomycin and removed from plates with Trypsin in EDTA. While the pigs were still
under the effect of anesthesia from the hysterectomy, 1.2x106 of the following human
tumor cell lines were injected: Panc01 (pancreas); U-251 (brain); HT-29 (colon); A549
(lung); or HepG2 (liver). Tumor cells were injected, subcutaneously, in 100 µL of
Matrigel behind the right ear, with an additional 100 µL of Matrigel behind the left ear.
Tumors were measured three times per week with plastic Vernier calipers. The diameter
was calculated as the square root of the product of the widest diameter and the
diameter perpendicular to that. At the same point of tumor measurement, weight and
other health monitoring was also completed. In addition to the subcutaneous cell line-
based tumors, we have also conducted orthotopic engraftment with human PDX tissues
and organoids [30, 51], significantly expanding the utility of these model animals.
4.3.5 Histotripsy Treatment of Porcine Pancreas and Liver
The ability to xenograft tumor cell lines and PDX tissue in the RAG2/IL2RG
knockout pigs orthotopically, provides a highly unique model for histotripsy
development. Our initial studies have focused on the pancreas, due in part to the high
relevance and difficulty in targeting that includes similar acoustic windows to that
expected in human patients. Initial studies have focused on refining the imaging and
targeting healthy organs with histotripsy in preparation for planned tumor ablation
studies in the RAG2/IL2RG knockout orthotopic pig model. Animals were anesthetized
109
with Telazol/Ketamine/Xylazine (TKX) and maintained under anesthesia for the duration
of the imaging and histotripsy treatment. Pigs were cleaned, dried, and had the area of
interest shaved and Nair (Naircare, Ewing, NJ, USA) was applied for 10 minutes before
being washed with water to thoroughly remove hair that could interfere with imaging.
Before treatment, freehand ultrasound imaging was used to identify the desired
treatment location using both a linear ultrasound imaging probe with a frequency range
of 10-18 MHz (L18-10L30H-4, Telemed, Lithuania, EU) and a 3 MHz curvilinear imaging
probe (Model C5-2, Analogic Corp., Peabody, MA). For treatment, the histotripsy
transducer and coaxial imaging probe were mounted onto a mechanical arm with an
associated micropositioning system to provide a uniformed volumetric ablation guided
by treatment planning software (Histosonics Inc., Ann Arbor, MI) [1]. The focus of the
transducer was aligned on the imaging screen before treatment by generating a bubble
cloud in degassed water. For treatment, the transducer was grossly positioned over the
pig’s abdomen and the focus was aligned to the targeted location using ultrasound
imaging. The transducer was coupled to the pig using a degassed water bath and an
adhesive drape, as shown in Figure 2C. Once this location was identified, the
transducer was turned on and histotripsy was applied to the target using single cycle
pulses and a PRF of 500Hz, with the pressure slowly increased until a bubble cloud was
generated in the tissue. The transducer was scanned to cover a 1 cm diameter
spherical ablation volume with 500 pulses applied per treatment location. Real-time
ultrasound imaging was used to monitor the treatment throughout the procedure. After
treatment, freehand ultrasound imaging was again conducted to assess for any tissue
damage, and the animals were then immediately euthanized per protocol by an
110
intracardiac injection of pentobarbital sodium and phenytoin sodium (Euthasol). A full
necropsy was conducted post-treatment by a board-certified veterinary pathologist
(S.C.O.).
4.3.6 Spontaneous Tumors in Canine Patients
In addition to the development of the SCID-like pig tumor model described above,
our team has recently initiated three studies testing histotripsy tumor ablation in
spontaneous tumors in client-owned dogs at the Animal Cancer & Research Center
(ACCRC), Virginia-Maryland College of Veterinary Medicine (VMCVM). These
prospective clinical trials evaluate the safety and feasibility of treating canine soft tissue
sarcoma and osteosarcoma tumors, which are currently underway, as well as a study
for treating canine brain tumors and others in future studies. These initial trials parallel
previous veterinary clinical trials at our site treating canine soft tissue sarcoma, liver,
brain, pancreatic, and lung cancers with either irreversible electroporation (IRE) or High
Intensity Focused Ultrasound (HIFU) thermal ablation. All of these canine tumor
populations represent potential candidates for future histotripsy studies. All clinical trials
are approved by the Institutional Animal Care and Use Committee and the Veterinary
Hospital Board. Client-owned dogs with naturally occurring tumors are recruited for
these clinical trials through the ACCRC referral network and by registry of the trial on a
publicly accessible, national veterinary clinical trials database [55]. All dogs undergo a
screening process to determine eligibility for trial enrollment, including a physical
examination, mass cytology or histopathology, complete blood count (CBC), serum
111
biochemistry profile, and thoracic and abdominal imaging. Standard-of-care treatment
options are always presented to owners and dogs are only included in studies if they
meet the inclusion/exclusion criteria and if a written owner’s informed consent is
obtained. Before a study treatment is administered for our current histotripsy clinical
trials, enrolled dogs will typically undergo cross-sectional imaging of the tumor either
with contrast-enhanced CT or MRI. These imaging studies enable visualization of the
tumor characteristics as well as the surrounding and overlying tissues to assess the
available acoustic window for each study. The planned histotripsy treatment is patient-
specific and scheduled as appropriate, with all further diagnostic evaluation and
treatments performed by the Histotripsy Clinical Team at the ACCRC.
4.3.7 Histotripsy Ablation of Spontaneous Canine Tumors
The feasibility of histotripsy tumor ablation in canine patients with spontaneously-
occurring tumors was tested in two dogs, one with a peripheral soft tissue sarcoma and
one with a hindlimb osteosarcoma tumor, using the 500 kHz clinic prototype histotripsy
system (Fig.2). Patient-specific histotripsy treatment plans were designed prior to each
treatment based on pre-treatment CT imaging, physical examinations, and ultrasound
imaging assessments. These measures were used to determine the acoustic window
and the prescribed area within the tumor that was targeted with histotripsy. For
treatment, the patients were placed under general anesthesia following standard
anesthetic protocols for client-owned patients. Anesthesia was maintained with inhaled
isoflurane. The fur was shaved as closely to the skin as possible over the planned
112
treatment site. Histotripsy was applied to 3cm diameter spherical volumes within the
tumor by sweeping the focal point across the target volume using the robotic micro-
positioner directed by the planning software. Vital signs (heart rate, blood pressure,
temperature, ECG, and SpO2) were monitored throughout the procedure to assess
patient safety and to identify any potential physiologic disturbances. To ensure acoustic
propagation from the transducer to the skin, a water bolus was coupled to the canine
patient over the optimal acoustic window identified using pre-treatment CT images and
freehand ultrasound imaging prior to treatment. The tumor was treated with histotripsy
under ultrasound guidance using single cycle pulses applied at a PRF of 500 Hz. Before
starting the volumetric treatment, the pressure at the focus was increased until a bubble
cloud was generated at the focus. A volumetric ablation volume was then generated
using the automated treatment strategy guided by the micropositioner and software of
the clinical histotripsy system. During histotripsy, the bubble cloud and tissue effects
were monitored using real-time US imaging. Upon completion of the treatment, patients
were recovered from anesthesia and were discharged into the care of their owners. The
patients received a physical examination and evaluation of the skin overlying the
histotripsy treatment site one day after treatment. Grading of adverse events was
determined according to the Veterinary Cooperative Oncology Group-Common
Terminology Criteria for Adverse Events (VCOG-CTCAE). A post-treatment CT scan
with contrast was performed one day after treatment to assess the completeness of the
ablation zone, the contrast enhancement pattern in the ablation zone, and the integrity
of any critical structures within or adjacent to the ablation zone. Surgical resection was
performed one day (osteosarcoma) or four days (soft tissue sarcoma) after histotripsy
113
treatment. A second post-treatment CT scan was performed immediately before
resection for the soft tissue sarcoma group. The patients were placed under general
anesthesia following standard anesthetic protocols for client-owned animals to facilitate
standard surgical resection of their tumors. The histotripsy-treated area of tumor and
bone (for osteosarcoma patients), and all relevant adjacent and overlying tissue were
inspected for gross signs of damage and harvested for histological evaluation. All study
procedures complied with the NIH Guide for the Care and Use of Laboratory Animals.
4.3.8 Histology & Morphological Analysis
After treatment, tissue specimens were visually inspected and fixed in 10% formalin
for at least 24 hours before sectioning and staining. All tissues were stained using
hematoxylin and eosin (H&E) to assess for histotripsy damage and any other changes
in structure and density of collagen and other tissue structures (i.e. vessels and ducts)
within the ablation volume. To determine the estimated extent of tissue ablated within
the targeted volume, individuals trained by a board-certified veterinary pathologist
compared regions of cellular damage with regions that were outside of the ablation
zone. Results were then verified by a veterinary pathologist (S.C.O.) to ensure accuracy
and avoid biases.
114
4.4 Results
4.4.1 Histotripsy Systems for Small and Large Animal Studies
Two separate histotripsy systems were used to evaluate histotripsy tumor ablation in
small (mice) and large (pigs, dogs) animal models. These separate systems were
needed to allow histotripsy to be precisely targeted to the tissues in these models based
on the tumor/tissue depths and acoustic windows available. In general, these in vivo
systems are representative of size, geometry, frequencies, and other properties of
transducers that have previously been used for applying histotripsy in small and large
animal models, respectively [2, 3, 12, 15, 20, 56].
For the small animal tumor ablation studies in the murine models, a miniature 1MHz
histotripsy transducer with a high frequency ultrasound imaging probe (Fig.1B) was
constructed to allow a well-confined bubble cloud to be generated within the shallow
subcutaneous or orthotopic tumors. This transducer was attached to a custom
motorized positioning system (Fig.1A) designed to apply automated volumetric ablations
closely mimicking those applied by the clinical histotripsy system. This transducer has a
full width half-maximum (FWHM) dimensions at a geometric focus of 0.98 mm, 0.93
mm, and 3.9 mm in transverse, elevational, and axial directions, respectively. Using this
system, a small, well-defined histotripsy bubble cloud can be generated at the focus
with approximate dimensions of ~2-3 mm in the axial directions and ~1-1.5 mm in the
transverse and elevational directions, depending on the pressure level applied. An
115
example high-speed image of the bubble cloud showing the approximate dimensions of
the cloud is shown in Figure 1E. Based on the working distance of the transducer, this
system is capable of treating shallow tumors in small animal models at depths up to ~1-
1.5 cm. The smaller bubble cloud generated by this small animal system, in comparison
to clinical histotripsy devices, also allows for more precise treatments and more fine
control over ablation margins, which is necessary in order to safely and effectively
generate histotripsy ablation of murine tumors without causing off target injury to
adjacent or overlying tissues. The treatments using this system are guided and
monitored by real-time ultrasound imaging using a high frequency linear ultrasound
imaging probe aligning co-axially inside of the therapy transducer. This higher frequency
imaging (10-18 MHz) provides superior imaging quality compared to the clinical imaging
systems commonly used in histotripsy procedures, including the large animal system in
this study, allowing for high resolution images of the superficial tumor tissue, the
histotripsy bubble cloud, and the resulting tissue damage generated by histotripsy.
For the large animal studies in both pigs and dogs, a custom 500 kHz transducer
was integrated onto a clinical histotripsy system provided to our team by HistoSonics
(Fig.2). This cart-based therapy system consists of a diagnostic ultrasound imaging
system embedded onto a cart with a mechanical articulating arm and micro-positioning
system that allows for precise volumetric ablations to be generated with histotripsy
(Fig.2A). A custom 500 kHz transducer was designed with a central hole to allow for the
system’s 3 MHz curvilinear imaging probe to be coaxially aligned inside of the therapy
transducer for real-time image guidance (Fig.2B). The therapy transducer was then
mounted onto the robotic micro-positioner in order to allow histotripsy to be uniformly
116
applied to a predetermined volume within the tissue, as outlined in a previous study
[15]. The transducer had FWHM dimensions at the geometric focus of 2.1 mm, 2.1 mm,
and 6.6 mm in transverse, elevational, and axial directions, respectively. Using this
system, a well-defined histotripsy bubble cloud could be generated at the focus with
approximate dimensions of ~5-6 mm in the axial directions and ~1.5-2.5 mm in the
transverse and elevational directions, depending on the pressure level applied during
treatment. An example high-speed image of a bubble cloud generated by the 500 kHz
transducer is shown in Figure 2E. Based on the working distance of the transducer, this
system is capable of treating tissue depths up to ~7 cm, allowing for treatments of the
pancreas and liver in pig models and a wide range of tumor types in canines including
superficial musculoskeletal tumors as well as abdominal cancers. Similar to the small
animal system described above, the histotripsy treatments applied using this large
animal device were all guided and monitored by real-time ultrasound imaging. However,
comparing the two systems, the large animal device provides lower resolution imaging
but more clinically relevant assessments of image-guidance for histotripsy devices that
are being developed for human applications by utilizing lower frequency imaging probes
to image at clinically relevant treatment depths in the pig and dog models that more
closely match human anatomy. Finally, it is worth noting that the larger bubble cloud
generated by the large animal histotripsy device allows for more rapid ablation of larger
treatment volumes using histotripsy while maintaining sufficient treatment precision for
targeting a predetermined volume within the target tissue.
117
4.4.2 Histotripsy Treatment of Subcutaneous Murine Tumors
Although a wide variety of subcutaneous and orthotopic tumor models have been
established in mice, the majority of histotripsy studies, including the example histotripsy
treatments shown in this study have focused on subcutaneous tumor injection models
that allow straightforward assessments of tumor progression and treatment with minimal
concerns regarding acoustic windows. In general, nodular tumors developed at the
injection site for all cell line and PDX based tumors. When studying cell line based
tumors in mice, it is well characterized that most tumors tend to be less physiologically
similar to the respective spontaneous tumors in human patients. For example, the
pancreatic Pan02 tumors are densely cellular with only minimal fibrovascular stroma
(Fig.3A). The tumors often have variably-sized foci of necrosis, which typically develop
centrally within the tumor. In terms of morphology, the tumor cells typically are arranged
in vague, indistinct streams. The cells often exhibit significant pleomorphism, including
differences in nuclear size and shape across the population with numerous mitotic
figures observed throughout cells within the tumor. This is significantly different
compared to a representative primary pancreatic tumor from a human patient (Fig.3B).
We have found human patient derived xenograft (PDX) tumors in NOD scid gamma
(NSG) mice to be highly useful in reproducing the complexity and stroma of the human
primary tumor, including for studies of pancreatic cancer [45]. The PDX models have
also proved useful for studies in multiple other cancers, including the prostate, liver,
breast, and colon, shown here as a model for cholangiocarcinoma (CC) (Fig. 3C). The
representative CC image shown in Figure 3C reflects the complex, multi-cellular tumor
118
with stroma and cells arranged into disorganized structures similar to de novo human
tumors.
Figure 4-3 Histotripsy Tumor Ablation in Murine Models. (A) Subcutaneous murine pancreatic Pan02 tumor. (B) Representative primary human pancreatic tumor. (C) Human CC PDX murine xenografted tumor. (D) Ultrasound images of histotripsy bubble clouds forming in RCC, (E) 4T1, and (F) CC PDX tumors. (G-I) Histology demonstrating full ablation within histotripsy targeted (G) RCC, (H) 4T1, and (I) CC PDX tumors.
Results from the histotripsy treatments of the subcutaneous RCC, PDX CC, and
orthotopic 4T1 mammary tumors showed that histotripsy was able to generate well-
defined bubble clouds within each tumor using our small animal system that were
clearly visualized on real-time ultrasound imaging (Fig. 3D-F). Using the automated
119
ablation strategy, volumetric ablations of the targeted regions of the tumor were
achieved for all three tumor types (Fig. 3G-I). While the central region of the tumor
targeted with histotripsy was fully ablated, the external margins of the tumor was still
intact after treatment, similar to previous reports [19, 20, 56]. More complete ablations
of the entire tumor volume could not be achieved in the small animal models due to the
inability to treat sufficient margins around the tumor, particularly on the portion of the
tumor located immediately below the skin. When treating near these regions, prefocal
cavitation was often noted during treatment at the skin-water interface, limiting the
ability to generate a bubble cloud immediately below the skin and causing potential
cavitation damage to the skin surface, which was observed in some subjects. With the
stiffer CC tumors, the bubble cloud appeared smaller and less dynamic on ultrasound
imaging compared to the other tumor types. For this reason, the CC treatments were
applied at a higher treatment dose with finer spacing between adjacent treatment points
in order to ensure complete ablation of the targeted tumor volume. It was also noted
that there was more variability in the appearance of the bubble cloud throughout
treatment in heterogeneous tumors, with regions of suppressed cavitation. These areas
of suppressed cavitation likely correspond to the more fibrous regions of the tumor.
Overall, results showed that histotripsy could be successfully applied to all murine tumor
types tested in this study, resulting in complete ablation of the targeted central region of
the tumor with the primary limitation being the inability to treat the entire tumor volume
and a clinically relevant treatment volume around the tumor. Regardless of the
limitations, we have found that the mouse models utilized thus far for histotripsy
development, including those shown in this study, are ideal for rapid and reproducible
120
parameter evaluation, hypothesis testing, and effective assessments of biological
hallmarks of cancer impacted by treatment. These models can also be used to assess
key engineering parameters necessary for device scale-up in certain cases, such as
comparing the relative differences in pulsing parameters needed to ablate different
tumor types.
4.4.3 Characterization of RAG2/IL2RG Pig Tumor Model
Our research team has previously generated and utilized immunodeficient pigs in a
variety of infectious disease and human xenograft tissue studies [57-59]. However, the
use of these pigs as cancer research models is still unique. As previously reported, we
utilized the CRISPR/Cas9 system to target the porcine RAG2 and IL2RG genes [57].
For the work described here, we utilized 454 in vitro fertilized oocytes, which were
injected with the targeted RNA for CRISPR/Cas9 gene disruption, followed by in vitro
culture [57]. Four days post-IVF, 155 embryos between the 8-cell and morula stage
were transferred to surrogate gilts. This resulted in 6 piglets being delivered by
hysterectomy into our germ-free house system, 114 days post-IVF. Genomic DNA was
collected from tail-snips for genotyping, which revealed that all 6 piglets were
successfully targeted for both the RAG2 and IL2RG genes, similar to our previous
reports [57]. Further functional verification from peripheral blood specimens, evaluated
via flow cytometry, confirmed that the piglets presented with the B-T-NK-SCID
phenotype.
121
Figure 4-4 RAG2/IL2RG Immunocompromised Pig Model for Cancer Research. (A) Example of a Pan01 (pancreatic cancer) subcutaneous tumor nodule growing behind the ear and (B) once excised. (C) Control Matrigel plugs exhibits no growth. (D, G) U-251 Human Glioblastoma, (E, H) 4T1 murine breast cancer, and (F, I) HepG2 human HCC are shown by histology under low and high magnification, respectively.
For tumor engraftment, we have evaluated tumor progression of the following human
cancer cell lines in the RAG2/IL2RG knockout pigs: PANC-1 (pancreatic cancer) (Fig.
4A-B); HepG2 (liver cancer); and U-251 (brain cancer). The murine 4T1 (breast cancer)
was also engrafted. The 6 piglets were randomized and were subcutaneously injected
with 1.2x106 cells in 100 µL of Matrigel in the ear. Control injections were also added,
which were 100 µL of Matrigel alone (Fig. 4C). Each cell line was injected in triplicate in
122
three separate pigs. In these initial studies, we established engraftment proficiency and
growth rates for each cell line. Within 24 hours, of the initial injection, we observed
tumor growth for all of the cell lines, except for 1, U-251 injection. This single U-251
injection was the only cell line that failed to engraft. The remaining cell lines rapidly
expanded over a 5-day period, eventually stabilizing and steadily increasing to the
target diameter of 1.0 – 1.6 cms by day 36. No changes in any clinical parameters were
noted for any of the animals. All tumors were readily palpated and observed by day 36,
whereas the Matrigel control injections were not detected. Tumors were evaluated by
ultrasound, revealing clearly defined tumors and clear delineations from surrounding
tissues. Extra-dermal, sub-dermal, and ultrasound tumor measurements were collected
at necropsy and compared for each tumor. Metastasis was also evaluated in the lung,
liver, spleen, and draining lymph nodes, with none detected. Histopathology
assessments were consistent with each tumor cell line, here showing U-251 (Fig. 4D
and G), 4T1 (Fig. 4E and H), and HepG2 (Fig. 4F and I) tumors as representative
specimens. Each was indistinguishable from the respective cell line based tumors
grown in immunocompromised mice. However, the size of the mice and the number of
tumors that can be generated is a significant limitation. This is especially true when
transitioning these studies to orthotopic tumor engraftment, where the mouse has
significant limitations in terms of human relevance. Future and on-going studies will
utilize these data to refine and optimize orthotopic tumor engraftment with these cell
lines in the RAG2/IL2RG deficient pigs. In addition to cell line tumors, we have also
successfully generated PDX-like tissue and tissue organoids from humans and
engrafted these more complex and physiologically relevant specimens into the
123
RAG2/IL2RG knockout animals, opening the opportunity to move beyond cell line-based
tumor models.
Figure 4-5 Histotripsy Treatment of Porcine Pancreas and Liver. (A) Image of the water bowl and site preparation with fluid repellent adhesive surgical drape for liquid containment during histotripsy. (B) Acoustic window for pancreas ultrasound treatment identified on pre-treatment imaging. (C) Gas in the stomach and GI tract encountered prior to histotripsy treatment. (D-E) Guided by real-time ultrasound imaging, histotripsy generated a clearly defined bubble cloud in (D) liver and (E) pancreas. (F) Gross image of 0.5 cm lesion in pancreas following histotripsy. (G) Off-target effects around the stomach and GI tract. (H-I) Representative histopathology images confirm regions of ablation in the pancreas.
124
4.4.4 Histotripsy Treatment of Porcine Pancreas and Liver
The feasibility of targeting the liver and pancreas in porcine subjects was tested
using the large animal histotripsy device shown in Figure 2. For this feasibility study,
pigs were imaged and treated with histotripsy in order to establish the regions of the
liver and pancreas that could be targeted for future orthotopic tumor studies in the pig
model described above. Treatments were applied non-invasively through an abdominal
window and partial ribcage obstruction (Fig.5A). The liver and pancreas were imaged
and targeted in each pig at treatment depths of ~4-6 cm (Fig. 5B-E). In the pancreas,
gas in the stomach and gastrointestinal (GI) tract was a significant issue (Fig. 5C). For
liver histotripsy treatments, well-defined bubble clouds were generated at the focus of
the transducer and clearly visualized as dynamically changing hyperechoic regions at
the focus of the transducer, as monitored by real-time ultrasound imaging (Fig.5D).
These results were consistent with previous in vivo hepatic histotripsy treatments that
have been reported in pig models [3, 11, 12, 15]. In contrast to the results seen in liver,
targeting the pancreas with histotripsy posed a more difficult challenge using our current
large animal system. Prior to treatment, the pancreas could be identified and visualized
on ultrasound imaging when using a freehand ultrasound imaging probe with significant
mechanical force applied to the abdomen to displace bowel gas (Fig. 5B-C). However,
the quality of the image was reduced significantly when the pancreas was imaged by
the co-axially aligned imaging probe within the therapy transducer (Fig.5E), likely due to
blockage from bowl gas (mechanical force couldn’t be applied in this case due to the
water standoff). In all subjects, the offset placement of the histotripsy transducer and
125
imaging probe through the degassed water coupling bowl prevented the pancreas from
being clearly visualized during treatment. Based on this limitation, the focus of the
histotripsy transducer was aligned to the region expected to contain the pancreas using
nearby anatomical landmarks, and histotripsy was then applied to this as described in
the Methods. During treatment, histotripsy cavitation could be heard, but the bubble
cloud was only sporadically visualized in the focal region during treatment (Fig. 5E). In
addition, prefocal cavitation was sporadically observed during treatment in regions
outside of the focus, likely corresponding to regions containing gas-filled overlying
tissues. After treatment, gross morphology showed a small histotripsy lesion in the
pancreas, as indicated by a region of localized hemorrhage approximately 0.5 cm in
size (Fig.5F). In addition to the damage to the pancreas, off target injury to surrounding
organs, including the small intestine and stomach, was observed (Fig.5G), likely
corresponding to the regions of prefocal cavitation observed during treatment. Ablation
in the pancreas was confirmed by histopathology as a small region of localized tissue
damage inside of the pancreas (Fig.5H-I).
4.4.5 Veterinary Clinical Oncology Patient Populations for Histotripsy Development
Moving beyond small and large animal cancer models, we have also utilized
histotripsy in the veterinary clinic with translational value to human clinical studies. The
Virginia-Maryland College of Veterinary Medicine (VMCVM) Animal Cancer Care &
Research Center (ACCRC) sees a large number of canine and feline oncology patients
with a wide range of pathologies. Patients are referred from general practice
126
veterinarians, veterinary specialists at external specialty hospitals, and internally from
clinical services within the VMCVM. Clinical trial participants are recruited from all
patients presenting to the ACCRC, supplemented through the Veterinary Clinical Trials
Network and Clinical Research Office (VCRO) in collaboration with the NCI
Comparative Oncology Trials Consortium (COTC)[37] and the NCI Comparative Brain
Tumor Consortium (CBTC)[60] to facilitate recruitment of patients for clinical trials. The
VCRO has a history of successfully recruiting companion animals for clinical trials and
has provided support for a number of canine and feline trials [61]. The following sections
describe the prevalence and characteristics of the most common malignancies seen at
our site, which is expected to be representative of other large veterinary hospitals in the
U.S.
In the veterinary clinic, soft tissue tumors are currently the most accessible tumor
types to treat using histotripsy and the data from subcutaneous tumors in animal models
can be directly applied. Tumors of the skin and subcutaneous tissues are common in
dogs, with malignant tumors comprising 20-40% of skin masses submitted for
histopathology [62]. Soft tissue sarcomas (STS) arise from the connective tissues,
comprising 15% of all skin tumors in dogs, and are subclassified pathologically based
on the tissue of origin [63]. Common tumor types seen in clinical practice include
fibrosarcoma, perivascular wall tumor, peripheral nerve sheath tumor, liposarcoma, and
myxosarcoma. Histologic distinction can be challenging and may be of minor clinical
importance as the majority of STS share a similar biological behavior and follow a
similar course of treatment: the patient could be cured with wide surgical excision, rarely
requiring adjuvant chemotherapy. The anatomical distribution varies, with most STS
127
occurring on the trunk and limbs. Generally, STS are slow-growing and not life-
threatening. However, in veterinary practice, this often results in medical attention being
sought late in the course of disease when the curative surgical treatment option may be
limb amputation, a disfiguring surgery, or a curative surgery is no longer possible. The
range of tumor sizes seen in typical veterinary specialty practice can range from a few
centimeters up to 20 cm. Our research team has had a significant degree of success
treating STS tumors using a range of tumor ablation modalities, including HIFU which
was applied in a recent study using the Theraclion Echopulse system which is available
at our site (Fig. 6A-D).
In addition to STS, many other tumor subtypes are of interest for histotripsy,
including bone tumors. Osteosarcoma (OS) is a primary bone cancer and a devastating
disease for both human and canine patients. It is the most common primary bone tumor
in children and adolescents, as well as in the dog [64, 65]. Figure 6F shows a CT image
of an osteosarcoma in the forelimb of a dog. Treatment of OS involves removal of the
primary tumor, and chemotherapy to inhibit metastatic disease. The main cause of
death in canine and human patients with OS remains metastatic disease.
Advancements in OS therapy need to focus on treating both primary and metastatic
tumors. Dogs and humans with OS share striking similarities that make the dog an
exceptional comparative oncology research model. These similarities include matching
biological behavior, nearly identical histological features, shared global gene expression
signatures, and similar responses to standard treatment regimens [66-70]. For example,
most OS lesions in dogs and humans occur in the appendicular skeleton around the
metaphyseal region of long bones, and the disease tends to be associated with large
128
and/or tall individuals [65, 66]. The anatomic similarities between the canine and human
skeletal structures allow the dog to be an accurate model for histotripsy systems
development and treatment evaluation studies. Osteosarcoma appears radiographically
identical in humans and dogs, displaying a lesion on CT and MRI scans with a
combination of lytic and proliferative bone [66, 71]. These radiographic similarities allow
for cross-species comparisons when evaluating histotripsy treatment outcomes using
imaging modalities.
The strength of naturally occurring canine OS as a model for comparative
oncology histotripsy research lies not only in the similarities of the disease in both
species but also in non-disease-related factors. The dog is an outbred species with an
intact immune system that develops spontaneously occurring OS over an extended
period, similar to humans and unlike inbred laboratory rodents. Dogs with OS reflect the
heterogeneity of human patients with OS, displaying interindividual variations in tumor
characteristics. Both humans and dogs with OS share similar clinical disease
manifestations and outcomes. The >10-fold increased incidence of canine OS
(estimated incidence of 13.9/100,000) compared to human OS (1.02/100,000), the
shorter lifespan of dogs, the rapid progression of metastatic OS in dogs enable
outcomes to be assessed in relatively short periods compared to their human
counterparts [66, 72, 73]. Metastatic bone tumors also occur in dogs, and common
primary cancers that metastasize to bone include mammary carcinoma, prostatic
carcinoma, urogenital carcinoma. Osteosarcoma can also metastasize to bone. The
ACCRC treats approximately 25 canine patients with OS annually, and this number is
expected to increase with recruitment for clinical trials. Histotripsy, with its ability to
129
disintegrate solid tumors and its potential for immunogenic stimulation, is uniquely
poised to have the potential to achieve the goals of OS therapy – treating both the
primary tumor and metastatic lesions.
Liver cancer is a preferred target for histotripsy and is the focus of the first human
clinical trial of the ablation strategy in patients [18]. Primary liver cancer in humans is
the fifth most common cause of cancer death in men and the ninth most common cause
of cancer death in women. Over the past two decades, incidence rates have increased,
and the overall mortality rates have been rising an average of 2.6 percent each year
[74]. At present, only 10-23% of human patients with HCC are surgical candidates for
curative-intent treatment [75, 76]. Liver transplantation is the only therapeutic option that
offers a consistent 90% survival rate, but the scarcity of liver donors severely limits its
broad applicability [77]. Primary liver tumors in dogs are considered infrequent and
account for 1.5% of all canine tumors, with hepatocellular carcinoma (HCC) being the
most common (Fig. 6G) [78]. The prognosis is good for massive HCC when surgical
resection is possible [79]. In contrast, the prognosis is poor for dogs with malignant
tumors other than massive HCC, and especially for dogs with nodular and diffuse HCC,
as surgical resection is not possible. Literature suggests that 40% of canine HCC are of
nodular or diffuse presentation and that a significant proportion of the massive HCC
tumors are non-resectable with complete margins due to their hilar or bile duct
association. Bland embolization and chemoembolization have been reported with
variable success in the palliation of four dogs with HCC [80].
130
Figure 4-6 Example Spontaneous Tumors in Canines. (A) Soft Tissue Sarcoma (STS) presented as a firm, fixed, painful soft tissue mass at the right proximal forelimb of a dog. (B) Contrast-enhanced CT consistent with a soft tissue attenuating and moderately contrast enhancing mass (yellow asterisk). The mass lies lateral to the triceps muscle and arises from the ascending pectoral muscle. (C) HIFU treatment of the tumor to partial ablation. An Echopulse unit by Theraclion was used. (D) Gross appearance of a cross-section of the tumor, 4 days post HIFU treatment. The yellow arrows point to the ablated tumor volume. (E) CT image of a rostral maxillary tumor in a dog, indicated in circled area. Note the aggressive invasion of the tumor into the maxilla, causing bone lysis. (F) CT image of a humeral osteosarcoma in a dog, indicated by the arrow. Note the aggressive nature of the tumor, causing bone lysis and proliferation. (G) CT image of a liver tumor in a dog, indicated by the circle. (H) CT image of a pancreatic tumor in a dog, indicated by the arrow. (I) CT image of a metastatic lymph node in the same patient, indicated by a circle.
131
In situ, energy-mediated tumor ablation is an alternative therapy to resection and
can be performed using a minimally invasive approach (percutaneous or laparoscopic)
[81]. Thermal ablation (radiofrequency and microwave ablation [RFA/MWA]) are the
mainstays of liver tumor ablation [81], with MWA currently superseding RFA as the
technology of choice [82]. Thermal ablation has similar oncological outcomes to
resection in carefully selected patients [83, 84]. However, it results in indiscriminate
tissue destruction at the near-field ablation zone, inducing a significant risk for
excessive, non-tumor tissue loss and damage to biliary and vascular structures,
resulting in high rates of liver-related complications [82, 85, 86]. Thermal HIFU has
enabled ablation of liver nodules and total tumor ablation is multiple studies for selected
human patients, but can also result in off-target effects to biliary and vascular structures
[87-89]. Thus, non-thermal ablative approaches are gaining interest in the treatment of
primary and metastatic liver tumors. Non-thermal ablative methods may also present a
unique opportunity to harness the patient’s immune system, as part of a multimodality
therapeutic approach. Rodent data suggest that tumor ablation by non-thermal
mechanisms can lead to anti-tumor immunity [90-92]. The liver represents an
immunologically rich and active organ, with intrahepatic immune cells skewed toward
tolerance, with quiescent or overtly suppressive functions [93, 94]. By attacking cancer
using non-thermal ablation, the tumor has the potential to serve as its own multivalent
anti-cancer vaccine through the presentation of tumor antigens and damage associated
signals originating from the ablated tissue [95]. At ACCRC, we have performed
approximately 15 HCC resections, about 8 liver tumor ablations annually, and we
132
evaluate an equal number of non-resectable cases. We demonstrated the feasibility and
safety of non-thermal liver tumor ablation using High-Frequency Irreversible
Electroporation (H-FIRE) by completing the first-in-patient pilot study of percutaneous
H-FIRE in dogs diagnosed with HCC. CT imaging before and 4 days after H-FIRE
indicated a predictable tumor ablation volume, and analysis of the treated tumor tissue
indicated specific T-cell recruitment and gene expression signatures associated with cell
injury/death and cell-mediated immunity [96]. Histotripsy represents an attractive, non-
invasive, non-thermal approach for the ablation of liver nodules. Histotripsy’s precise
ablation targeting capabilities in combination with the completely non-invasive approach
and the ACCRC’s experience in minimally invasive ablative approaches for liver cancer
would allow for the canine cancer patients diagnosed with liver cancer to serve as the
translational model for its application as an outpatient procedure.
Similar to humans, pancreatic cancer in dogs is also a critically important
malignancy. Pancreatic tumors in dogs can originate from the exocrine pancreas, for
example, adenocarcinomas of the pancreatic ducts or acini, or originate from the
endocrine pancreas, for example, insulinoma. Insulinoma is the most common
pancreatic endocrine tumor in dogs, originates from pancreatic cells and secretes
insulin, causing potentially life-threatening hypoglycemia [97]. Insulinoma also
aggressively metastasizes to other organs, commonly to the liver and regional lymph
nodes. Figure 6H-I shows CT images of a pancreatic tumor with its associated lymph
node metastasis in a dog. Canine insulinomas behave in a biologically similar fashion to
human malignant insulinoma [98]. The functional insulin-secreting nature of primary and
133
metastatic insulinoma is a major life-limiting factor for patients, and treatment of both
primary and metastatic disease is essential for improving prognosis.
Surgical resection of all gross disease in dogs with pancreatic cancer, though not
curative, is recommended to improve the efficacy of medical adjuvant treatments and
disease outcomes [97]. Complete surgical resection of all gross disease is not always
achievable, depending on the location and extent of the lesions. Resection of tumors
located in the body of the pancreas is associated with high morbidity and potentially
mortality rates, and the extent of metastatic lesions may preclude surgical resection.
Novel treatment approaches are needed to improve the long-term survival of pancreatic
cancer patients. New treatment techniques that can non-surgically ablate tumor lesions
are attractive alternates to surgical tumor resection. Treatment approaches to stimulate
the host immune system to mount an anti-tumor immune response that carries immune
memory will improve the prognosis for the development or recurrence of metastatic
disease in pancreatic cancers. Histotripsy, with its ability to precisely ablate solid tumors
and its potential for immunogenic stimulation, is uniquely poised to advance the
therapeutic efficacy and prognosis of insulinomas, a tumor with currently limited
treatment options. The ACCRC treats approximately 8 canine patients with insulinoma
annually, and this number is expected to increase with recruitment for clinical trials. The
ACCRC has treated insulinoma tumors in dogs using another investigational non-
thermal tumor ablation technique, H-FIRE, and will continue its investigations into
pancreatic tumor ablations using histotripsy.
Canine oral tumors are another tumor type that can serve as strong comparative
oncology research models for their human counterparts. For instance, canine oral
134
melanoma has been shown to represent a faithful model for human mucosal melanoma
and human triple wild-type melanoma [99-101]. The disease in both species shares
genetic signatures, clinical behavior, and histological characteristics [102, 103]. Head
and neck squamous cell carcinomas are the most common oral cancer in people, and
canine oral squamous cell carcinoma shares similarities with the human form of this
disease [104]. Patients with oral cancers commonly present with clinical signs
associated with pain due to the oral cancer, decreased appetite, and weight loss. Oral
tumors in dogs occur with relatively low frequency, constituting 6% of all canine
cancers, and three of the most common oral cancers in dogs are malignant melanoma,
squamous cell carcinoma, and fibrosarcoma [105-107]. Oral osteosarcoma (OS)
represents 12 - 13% of all canine OS [108]. Canine oral tumors are commonly locally
invasive and many are aggressively metastatic, especially to the locoregional lymph
nodes. A non-surgical tumor ablation technique such as histotripsy, which is also
capable of stimulating an anti-tumor immune response and can be used for tissue
selective ablation of tumors near critical structures, has exciting potential to benefit both
canine and human patients suffering from oral cancers. The ACCRC treats
approximately 8-10 canine oral tumors annually. Figure 6E shows an example of a
canine oral tumor that is locally invasive, causing bone lysis of the rostral maxilla.
Our research team has been highly successful in utilizing tumor ablation
modalities to treat canine brain tumors [109]. Despite significant progress over the last
decade, brain tumors such as malignant gliomas (MG), represent some of the most
treatment-refractory cancers in both humans and dogs, and local treatment failures
remain a significant source of morbidity and mortality in brain tumor patients. Dogs and
135
humans are the only mammals known to spontaneously and commonly develop MG,
and MG accounts for approximately 35-40% of all primary brain tumors in dogs. As
dogs have a complex gyrencephalic brain, and canine gliomas share many clinical,
imaging, histomorphologic, molecular, and genetic features with human gliomas, dogs
with brain tumors are uniquely suited translational model systems for the study of tumor
ablation strategies [90].
In both dogs and humans, there is a dire need for new approaches that can be
applied to tumors involving eloquent brain regions, circumvent or disrupt the blood brain
barrier, target highly resistant tumors and cancer stem cells, generate minimal collateral
damage to healthy tissue, and promote an anti-tumor immune response. Successful
therapeutic strategies are likely to require a combination approach involving surgery,
chemo and radiation therapy, and novel ablation approaches that can de-bulk the
primary tumor and attenuate recurrence driven from residual cancer stem cells
repopulating the resected tissue, or predictably and reversibly disrupt the blood brain
barrier in a penumbra of tissue surrounding the bulk tumor to facilitate the delivery of
therapeutics that are normally impermeant to the brain [110]. Histotripsy has been
utilized in pre-clinical porcine models to ablate brain tissue. However, it is still in the
early stages of development for the treatment of brain tumors in the veterinary clinic.
Unlike many of the other tumors discussed thus far, a significant challenge to the
application of histotripsy in the brain is the thickness inhomogeneities inherent to the
skull, which can cause aberrations in transcranially propagated ultrasound pulses
resulting in reduced focal pressure amplitudes in the target tissue. Thus, a craniectomy
is required to provide an acoustic window for effective bubble cloud formation. To
136
overcome current obstacles associated with histotripsy in the brain, our research team
has developed novel acoustically transparent cranial implants and is rapidly adapting
clinical histotripsy systems for use in active clinical trials in dogs with brain tumors.
4.4.6 Histotripsy Ablation of Spontaneous Canine Tumors
Figure 4-7 Histotripsy Treatment of Canine Soft Tissue Sarcoma. (A-B) Image of spontaneous soft tissue sarcoma with estimated tumor size of ~7-8 cm in diameter. (C) Contrast-enhanced CT collected before histotripsy treatment. (D) Histotripsy treatment experimental set-up. (E) Real-time ultrasound image collected during histotripsy treatment showing the bubble cloud visible as a hyperechoic, oscillating zone (arrow). (F) Contrast-enhanced CT collected 4 days after histotripsy treatment with decreased contrast uptake visible in the zone of ablation. (G) Gross morphological analysis of resected canine soft tissue sarcoma. Box indicates treatment region, identified by location and presence of tissue necrosis and hemorrhage. (H) H&E image of untreated canine soft tissue sarcoma (20x magnification) with intact cells and undisturbed extracellular matrix is observable. (I) H&E image of canine soft tissue sarcoma tissue treated with histotripsy (40x magnification) showing extensive necrosis and hemorrhagic appearance.
137
Figure 4-8 Histotripsy Treatment of Canine Osteosarcoma. (A-B) Image of an undisturbed spontaneous canine osteosarcoma in the left hind limb. Significant soft tissue swelling was present, creating an estimated zone of ~30 cm of affected tissue. (C) Contrast-enhanced CT collected before histotripsy treatment. (D) Histotripsy treatment experimental set-up. (E) Real-time ultrasound image collected during histotripsy treatment with the bubble cloud visible as a hyperechoic, oscillating zone (arrow). (F) Contrast-enhanced CT collected 1 day after histotripsy treatment.
The feasibility of conducting in vivo histotripsy treatments in canine patients with
spontaneous tumors was tested in two initial patients with a peripheral soft tissue
sarcoma (Fig.7) and a hindlimb osteosarcoma tumor (Fig.8). Histotripsy was applied
non-invasively at a p- estimated to be ~30-35 MPa. A 3 cm spherical ablation volume
was targeted inside of both tumors, and histotripsy was applied uniformly over this
volume at a PRF of 500 Hz and a dose of 500 pulses/point, with a total treatment time
of ~59 minutes. In both dogs, well-defined histotripsy bubble clouds were generated at
the focus of the transducer and clearly visualized as dynamically changing hyperechoic
regions throughout the treatment, as monitored by real-time ultrasound imaging (Fig.7E,
138
Fig.8E). The procedures in both dogs were tolerated well throughout treatment, with no
signs of distress or changes in vital signs during the procedures. After treatment, both
subjects were recovered until follow-up CT imaging and surgical resection one day
(osteosarcoma) or 4 days (soft tissue sarcoma) after treatment. Both the pre- and post-
treatment CT images (Fig.8C/F) for the dog with the osteosarcoma tumor did not show
a clearly defined tumor or ablation zone after treatment, likely due to the heterogeneous
nature of the osteosarcoma tumor, the presence of significant edema, or CT imaging
parameters. Future work will aim to improve the CT imaging for these patients or utilize
MRI imaging, as needed. The post-treatment CT image for the dog with the peripheral
soft tissue sarcoma showed a clearly defined ablation zone closely matching the region
targeted with histotripsy (Fig.7F). The ablation zone can be clearly visualized on CT as
a darkened region of the tumor cross-sectional image (Fig.7F), indicating a decrease in
uptake of contrast and blood flow to this region which correlates with the successful
histotripsy treatment. After treatment, gross morphology showed a histotripsy lesion was
generated in the targeted tumors, and the tissue was processed for histological analysis
after resection. Figure 7G-I shows an example of gross morphology and histology for
the peripheral soft tissue sarcoma tumor treated with histotripsy, with results showing
histotripsy generated complete ablation of the targeted tumor tissue. Together, results
from these initial treatments provide an example of histotripsy tumor ablation in dogs
with spontaneous tumors and demonstrate the potential of utilizing veterinary oncology
populations for conducting more clinically relevant studies of histotripsy for cancer
applications.
139
4.5 Discussion
While early diagnosis and multi-scale combinatorial treatment approaches are
essential to reducing cancer related mortality, the lack of treatment options for many
types of cancers has resulted in an unacceptable stagnation in patient morbidity and
survival. New treatment options and therapeutic strategies are critical to improve the
survival of these patients. Focal tumor ablation modalities have shown recent success
in multiple clinical trials by crossing barriers that are difficult to bridge by chemotherapy
and surgery. Histotripsy is emerging as one of the more promising modalities. However,
as with all of the other tumor ablation strategies, clinical translation for many of the most
difficult to treat malignancies has been hampered by the lack of robust pre-clinical
animal models. While some modalities, such as radiotherapy and microwave ablation,
have the historical advantage of multiple years of clinical data in human patients, it is
much more difficult for emerging technologies, such as histotripsy or IRE, to be gain
widespread adoption in the human clinic without more mechanistic insight that defines
their therapeutic advantages. Indeed, the non-thermal nature of these modalities
appears to have significant benefits and increasingly nuanced mechanisms that require
more robust assessments to fully understand their mechanisms of action. For example,
it is becoming increasingly apparent that these modalities are significantly better than
currently adopted strategies at engaging the immune system following treatment [19,
92]. This has significant implications associated with the so-called “abscopal effect”
[111], which appears to occur more predictably and robustly following non-thermal
ablation. This has multiple benefits, including reduced metastatic burden, lower
140
recurrence rates, improved responsiveness to immunotherapeutics, and improved
overall survival. However, the improved systemic effects associated with local tumor
ablation significantly increases the complexity of the studies necessary to validate these
mechanisms. Thus, necessitating the development of improved pre-clinical animal
models.
In terms of animal models, the use of rodents in tumor ablation studies continues
to be widely accepted. This is certainly understandable due to the historic use of mice
and rats in the cancer biology and biomedical engineering fields. Indeed, we have
already mentioned several advantages and disadvantages, such as the small size that
can be highly detrimental for engineering and device design. This often necessitates the
miniaturization of devices that work well in rodents but either fail to scale up and
translate well to humans or require additional validation in large animal studies prior to
regulatory approval for human testing. For example, the histotripsy system used to treat
the mouse tumors in this study consisted of a highly customized therapy transducer that
could only target small, superficial tumors at depths <1.5 cm. Using this system to treat
tumors in mice does not answer key questions about the potential feasibility of treating
tumors in human patients through the acoustic windows available for a given pathology
or the optimal approaches for achieving complete volumetric ablation of human tumors
(with adequate clinical margins) that are located in different regions and with a wide
range of characteristics including tumor size, the number of nodules, and the presence
of critical structures within or near the tumor. Furthermore, in addition to the limitations
of the therapy transducer, the small animal system also utilizes a high frequency
ultrasound imaging system that provides excellent real-time treatment guidance and
141
monitoring but only provides minimal information about the potential to guide and
monitor histotripsy treatments in future clinical studies. Although small animal tumor
studies are often supplemented with large animal experiments in healthy animals, such
as pigs, studies of histotripsy ablation of healthy tissue do not allow for investigations
into the effectiveness of histotripsy to achieve precise and complete ablation of a target
tumor volume nor the ability to assess real-time and post-treatment imaging methods to
separately visualize the ablation zone from residual untreated tumor. Finally, these
limitations make it difficult to assess the safety of histotripsy for treating larger tumors
that are typically seen in humans as well as the safety of ablating tumors in an animal
with similar anatomy and physiology to human patients.
In many ways, this failure to translate new ablation methods, such as histotripsy,
into the clinic is associated with the choice of cancer models used to evaluate, validate,
and optimize therapeutic strategies. For example, most tumor ablation studies have
taken advantage of subcutaneous tumor engraftment models, especially during early
modality development. As discussed in our results section, these models have several
advantages and are highly useful, especially during early proof-of-concept/proof-of-
principle testing. The subcutaneous engraftment of human cancer cell lines in
immunocompromised mice, for example in the flank, have a long history of use in
studying cancer hallmarks. In terms of histotripsy, these models offer a readily
accessible tumor where the direct effects of ablation can be evaluated, both visually and
by ultrasound. Likewise, the use of human cells in the mouse allows for straightforward
visualization of ablation using human and mouse-specific antibodies and other reagents
to label the human-derived tumor cells and host specific mouse cells. Unfortunately, as
142
shown and discussed in our results, tumors generated subcutaneously and using cell
lines have several disadvantages. Specifically, the tumor morphology, biology, and
stroma are significantly altered from the original host tumor. In the context of histotripsy,
this is a significant concern where tissue characteristics, particularly tissue fibrosis and
other factors that impact the local tissue mechanical strength, can significantly impact
ablation success [6, 14, 112-114]. Without adequate models that can recapitulate the
relevant tumor microenvironment, it is difficult to identify the optimal histotripsy
parameters and treatment doses needed to achieve complete tumor ablation or to
develop improved histotripsy strategies for developing tumor-selective ablation methods
that can ablate the tumors while preserving critical structures such as vessels, nerves,
and bile ducts [1, 3, 9, 13, 14]. Likewise, the lack of a functional immune system does
not allow the evaluation of this increasingly important aspect of the tumor
microenvironment that may have significant implications in histotripsy effectiveness.
The use of rodent derived cancer cells in the host species is a significant
improvement over the use of human cell lines in immunocompromised mice. These cell
lines are more amenable to orthotopic studies, where the cells can be engrafted into the
tissue of relevance in immunocompetent mice. This provides significantly increased
relevance, a more accurate host tissue stroma, and the intact immune system allows for
assessments of both local and systemic anti-tumor responses following treatment,
which is becoming more essential based on recent studies showing the potential
immunological benefits of histotripsy. The ability to genetically modify both the host and
the cancer cell line also allows for robust mechanistic studies. It should be noted that
the mouse strain and sex are both critical factors to consider in these models. For
143
engraftment to be successful, the strain (i.e. C567Bl/6 or BALB/c) of the host and the
cell line must match or the host immune system will reject the tumor. Likewise, male
derived tumor cells can fail to engraft in female hosts due to Histocompatibility Y antigen
incompatibility and tissue rejection. Thus, while the intact immune system is an
advantage, this can limit tumor models in rodents. Likewise, similar to the limitations
discussed above for human cell lines and as shown in our results here, the same
limitations associated with unrepresentative tumor microenvironments, lack of cell
diversity within the tumor, and inaccurate tumor-associated stroma all exist in mouse
cancer cell line derived tumors. Each of these are major issues for histotripsy
development.
To circumvent many of these limitations, our research team has recently
deployed PDX models for histotripsy development [45]. PDX rodent models utilize the
engraftment of cancer tissue, typically biopsy or surgically resected tissue fragments,
into immunocompromised mice. These mice are typically NOD scid gamma (NSG)
mice. As shown and discussed in our results section, over time these tumor fragments
will proliferate into a tumor that closely mimics the biological and stromal complexity of
the original patient’s tumor. The tumor can be expanded to provide ample tissue for ex
vivo/in vitro studies necessary for device development, the tissue can also be evaluated
in the animal. Likewise, the added ability to detect human cells in mice can provide high
resolution assessments of tumor ablation. While this method still suffers from the lack of
an immune system niche, this strategy is incredibly useful for the evaluation of tumor
specific ablation to define parameters that are impacted by unique tumor
microenvironments, cell complexity, and stroma characteristics.
144
Moving beyond mice, large mammal pre-clinical models can also be effectively
incorporated into tumor modality development. Here, we propose the incorporation of
pigs due to their overall anatomical and physiological similarities to humans. Likewise,
as we discuss in the results above, genetic manipulation is now becoming more
commonplace and the pig is emerging as a key cancer model due to the development
of the immunocompromised RAG2/IL2RG deletion animals that are receptive to cancer
cell engraftment and the rise of genetically modified animals, such as the “oncopig” [29].
In terms of histotripsy development, we have focused on the RAG2/IL2RG deletion pigs.
These animals are functionally similar to the NSG mice, mentioned above, and we have
found that they can be used in identical applications. Advantages of this model include
the ability to surgically implant human (or other species) cancer cell lines in specific
locations within organs of interest, such as adjacent to major blood vessels or nerves.
The cell lines have a predictable growth rate when implanted either subcutaneously or
in situ, allowing for a highly reproducible and effective model system to evaluate tumor
ablation modalities. It should be noted that the use of the CRISPR/Cas9 system allows
for the generation of animals “on-demand”, without the need for maintaining breeding
colonies. However, one disadvantage is that the targeting of the RAG2/IL2RG genes
are less predictable. For example, as mentioned in the results section, one of the U-251
cell lines did not successfully engraft in the pigs. Further genotyping of this animal
revealed only a partial knockdown of the RAG2 and IL2RG genes, resulting in an
attenuated but still viable immune system that drove tumor rejection. Thus, before
orthotopic injection, it is critical to fully determine both genotype and immune system
phenotype in these animals. It is also important to note that due to their
145
immunocompromised status, these animals require housing under germfree conditions
that are only available at a small number of institutions worldwide. Moving beyond
human cell line studies in this unique model, which have some of the same limitations
discussed above for the mouse studies, these animals are uniquely amenable to more
complex human tissue engraftments, such as from PDX tissues and organoids. Thus,
as we move into the future, these highly useful specimens will allow for orthotopic
engraftment in these unique animals for more robust histotripsy development,
refinement, and treatment optimization.
We have successfully incorporated porcine models into our tumor ablation
modality testing pipeline. In addition to the tumor models discussed above, healthy pig
models have already proven useful in defining histotripsy treatment parameters utilized
in current and planned clinical trials [15, 18]. For example, as we report here, we have
successfully refined the use of histotripsy in targeting the healthy liver in pigs. By
expanding these studies to include liver tumors in the pig models, we expect to take
advantage of the features of healthy pig models that are anatomically similar to humans
while also being able to compare tumor-specific therapy and imaging methods that can’t
be tested in healthy animals, such as the safety and effectiveness of ablating liver
tumors located near critical structures or the ability to visualize and target liver tumors in
locations that require acoustic access through partial or complete rib coverage. In
addition to the liver, the work in this study has started to extend the use of histotripsy as
a non-invasive method for ablating pancreatic tumors. In the results presented above,
we report here for the first time histotripsy being used to target the pancreas in a large
animal model. This is an important milestone, but also illustrates some of the difficulties
146
in treating the pancreas and further supports the use of large animal models, such as
the pig, in device development. The acoustic window is critical to most applications of
histotripsy, including pancreatic cancer, and it cannot be effectively replicated in mouse
models. As we report here, gas in the stomach and GI tract can significantly limit the
ability to visualize the pancreas during histotripsy procedures and may also limit
histotripsy from generating a precise and effective bubble cloud in the pancreas without
causing off-target cavitation injury. We have recently developed highly effective
protocols to better alleviate these issues, which range from engineering changes to the
device to refining our imaging and treatment parameters for use in pancreatic cancer,
as well as, investigating other changes such as diet adjustments, pharmaceutical
interventions, and surgical/interventional strategies to remove excess gas prior to
treatment. For example, we have successfully utilized the addition of custard to the pigs’
diet and optimized a cocktail of simethicone (anti-gas) and Bisacodyl (laxative) as a less
invasive strategy to improve the acoustic window, similar to studies reported in human
patients preparing for HIFU treatment [115, 116]. Ultimately, our histotripsy treatment
parameters will be combined with our tumor models described above. Consistent with
ongoing human and veterinary clinical studies, we anticipate that tumors in the
pancreas will be easier to image and target compared to the healthy pancreas. Moving
beyond the pancreas, in this report, we describe the use of multiple cell lines from
cancers currently being evaluated by our research team. In the future, we plan to
incorporate histotripsy data generated here from the liver and pancreas to refine the
treatment parameters of these other cancers and tissues, defined here in subcutaneous
models, including the use of PDX specimens and organoids to provide an even more
147
robust and translational model for both human and veterinary patients with a wide range
of cancer types.
In addition to human applications, tumor ablation modalities such as histotripsy
are also emerging therapeutic strategies in the veterinary clinic, especially in veterinary
oncology. Ablation techniques such as histotripsy offer an attractive alternative
treatment to surgical resection of tumors. The current standard-of-care for many tumors
in veterinary oncology includes surgical resection of the primary tumor. However,
surgical resection sometimes is accompanied by excessive risk and morbidity. For
example, limb salvage surgery in dogs to resect the primary tumor in OS is associated
with a high rate of complications, with major complication rates as high as 71% reported
[117]. Additionally, despite the advancements in chemotherapeutics, many veterinary
cancer patients, like their human counterparts, eventually succumb to metastatic
disease. Histotripsy, with its ability to disintegrate solid tumors and its potential for
immunogenic induction of an anti-tumor response, is uniquely poised to achieve the
major goals of cancer therapy – treating both the primary tumor and metastatic lesions.
Not only can veterinary cancer patients directly benefit from novel ablation technologies
such as histotripsy, they also provide excellent clinical trial populations for comparative
oncology studies to benefit humans. The anatomic, physiologic, and cancer biology
similarities between many of the animal species treated in the veterinary clinic to human
cancer patients offer a direct translation of technologies from bench-to kennel-to
bedside [118]. For example, the results from the histotripsy treatments of canine STS
and OS shown in this study highlight many of the benefits of testing histotripsy in dogs
with spontaneous tumors. More specifically, treatments were conducted with a clinical
148
prototype histotripsy system that consists of the therapy transducer, imaging system,
and volumetric ablation treatment strategies that are similar to those expected to be
used in humans. The results from these studies, which demonstrated histotripsy could
achieve precise and complete ablation of targeted volumes within both the STS and OS
tumors, are expected to be more directly translatable to humans when it comes to the
devices and treatment parameters used for histotripsy. In addition to providing a more
clinically relevant population for testing the safety and efficacy of histotripsy devices, the
dog as a comparative oncology research model also offers the unique advantage of
being an outbred species with an intact immune system that develops spontaneously
occurring tumors over an extended time, just as humans do, unlike inbred laboratory
rodents that are not syngeneic host-tumor models. Utilizing canine models of
spontaneous disease allows for evaluation of immune responses to treatments, an
evaluation that is not as readily achieved in immunodeficient rodent models. Dogs and
humans are more alike anatomically and physiologically than are mice to humans and
share the same physical environments, unlike laboratory rodents maintained in a highly
controlled research facility environment [70]. Dogs with spontaneously occurring tumors
represent powerful tools for device development and clinical trials, as they are a
biologically relevant disease model that can serve to better bridge preclinical murine
study findings with human clinical trials. Additionally, the willingness of canine owners to
engage in sample collections and procedures such as biopsies make dogs an
invaluable model for clinical testing. Conversely, client-owned dogs present limitations
such as the low prevalence of certain tumor types, for example, intestinal cancer,
compared to humans, thus limiting the study of these diseases.
149
4.6 Conclusion
Histotripsy is rapidly emerging as the first completely non-invasive, non-thermal, and
non-ionizing ablation method for the treatment of cancer. However, the lack of clinically
and physiologically relevant pre-clinical cancer models has slowed the translation of
histotripsy into human clinical trials. This paper highlights the pros and cons of a diverse
range of small and large animal tumor models, including spontaneous tumors in
veterinary patients that are currently being developed for investigating histotripsy for
oncology applications. Results of example histotripsy treatments performed in mouse
tumor models demonstrate the feasibility of using mice as highly cost-effective models
that can test histotripsy for a nearly limitless range of cancer types. However, results
also show the limitations of mouse tumor models due to their small size, lack of clinical
and physiological relevance to humans, and the inability to test clinically relevant
histotripsy devices and treatment protocols. To address these limitations, our team has
developed a novel RAG2/IL2RG deletion pig tumor model that is being developed for
histotripsy studies. Results from preliminary studies using this model show its potential
to be used with a wide range of tumor types to study histotripsy tumor ablation in a
clinically relevant large animal model. Preliminary histotripsy experiments in the healthy
pig liver and pancreas further demonstrated the strengths of this model, which will be
utilized in planned studies testing histotripsy pancreatic tumor ablation using orthotopic
tumors grown in the pig pancreas as well as future studies with additional cancer types.
Finally, this work highlights the potential benefits of developing histotripsy for the
treatment of spontaneous tumors in veterinary patients and utilizing these veterinary
150
patient populations to provide more realistic models for developing and testing
histotripsy devices and techniques designed for future use in human patients. Together,
these studies demonstrate the potential advantages of bench-to kennel-to-bedside
approaches in the development of histotripsy for oncology applications that should be
considered to improve the rate and success of efforts to translate histotripsy into clinical
practice for the treatment of cancer.
4.7 Acknowledgment
This work was supported by grants from the Focused Ultrasound Foundation (FUSF-
RAP-S-18-00011, FUS746), The National Institutes of Health (R21OD027062,
R21EB028429, R21EB030182, R21EB027979), Grayton Friedlander Memorial Fund,
and the American Kennel Club Canine Health Foundation. This work was also
supported by the Virginia Maryland College of Veterinary Medicine; The Virginia Tech
Institute for Critical Technology and Applied Sciences Center for Engineered Health;
and the Virginia Tech Department of Biomedical Engineering and Mechanics.
151
4.8 References
[1] C. M. Smolock AR, Vlaisavljevich E, Gendron-Fitzpatrick A, Green C, Ziemlewicz
TJ and Lee FT, "Robotically Assisted Sonic Therapy as a Non-Invasive Non-
Thermal Ablation Modality: Proof of Concept in a Porcine Liver Model,"
Radiology, vol. 287, no. 2, pp. 485-93, 2018.
[2] E. Vlaisavljevich et al., "Non-Invasive Ultrasound Liver Ablation Using Histotripsy:
Chronic Study in an In Vivo Rodent Model," Ultrasound Med Biol, vol. 42, no. 8,
pp. 1890-902, Aug 2016, doi: 10.1016/j.ultrasmedbio.2016.03.018.
[3] E. Vlaisavljevich et al., "Image-guided non-invasive ultrasound liver ablation
using histotripsy: feasibility study in an in vivo porcine model," Ultrasound Med
Biol, vol. 39, no. 8, pp. 1398-409, Aug 2013, doi:
10.1016/j.ultrasmedbio.2013.02.005.
[4] T. Worlikar et al., "Histotripsy for Non-Invasive Ablation of Hepatocellular
Carcinoma (HCC) Tumor in a Subcutaneous Xenograft Murine Model," Conf Proc
IEEE Eng Med Biol Soc, vol. 2018, pp. 6064-6067, Jul 2018, doi:
10.1109/EMBC.2018.8513650.
[5] A. D. Maxwell, C. A. Cain, T. L. Hall, J. B. Fowlkes, and Z. Xu, "Probability of
cavitation for single ultrasound pulses applied to tissues and tissue-mimicking
materials," Ultrasound Med Biol, vol. 39, no. 3, pp. 449-65, Mar 2013, doi:
10.1016/j.ultrasmedbio.2012.09.004.
152
[6] E. Vlaisavljevich et al., "Effects of ultrasound frequency and tissue stiffness on
the histotripsy intrinsic threshold for cavitation," Ultrasound Med Biol, vol. 41, no.
6, pp. 1651-67, Jun 2015, doi: 10.1016/j.ultrasmedbio.2015.01.028.
[7] Z. Xu et al., "Controlled ultrasound tissue erosion," IEEE Trans Ultrason
Ferroelectr Freq Control, vol. 51, no. 6, pp. 726-36, Jun 2004, doi:
10.1109/tuffc.2004.1308731.
[8] R. M. Xu Z, Hall TL, Mycek M-A, Fowlkes JB and Cain CA, "Evolution of bubble
clouds produced in pulsed cavitational ultrasound therapy - histotripsy," IEEE
Trans Ultrason Ferroelectr Freq Control, vol. 55, pp. 1122-32, 2008.
[9] E. Vlaisavljevich, A. Maxwell, M. Warnez, E. Johnsen, C. A. Cain, and Z. Xu,
"Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
mechanical properties," IEEE Trans Ultrason Ferroelectr Freq Control, vol. 61,
no. 2, pp. 341-52, Feb 2014, doi: 10.1109/TUFFC.2014.6722618.
[10] E. Vlaisavljevich, A. Maxwell, L. Mancia, E. Johnsen, C. Cain, and Z. Xu,
"Visualizing the Histotripsy Process: Bubble Cloud-Cancer Cell Interactions in a
Tissue-Mimicking Environment," Ultrasound Med Biol, vol. 42, no. 10, pp. 2466-
77, Oct 2016, doi: 10.1016/j.ultrasmedbio.2016.05.018.
[11] V. E. Kim Y, Owens GE, Allen SP, Cain CA and Xu Z, "In vivo transcostal
histotripsy therapy without aberration correction," Phys Med Biol, vol. 59, pp.
2553-68, 2014.
[12] E. Vlaisavljevich et al., "Non-Invasive Liver Ablation Using Histotripsy: Preclinical
Safety Study in an In Vivo Porcine Model," Ultrasound Med Biol, vol. 43, no. 6,
pp. 1237-1251, Jun 2017, doi: 10.1016/j.ultrasmedbio.2017.01.016.
153
[13] A. M. Lake, Z. Xu, J. E. Wilkinson, C. A. Cain, and W. W. Roberts, "Renal
ablation by histotripsy--does it spare the collecting system?," J Urol, vol. 179, no.
3, pp. 1150-4, Mar 2008, doi: 10.1016/j.juro.2007.10.033.
[14] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," (in eng), Phys Med Biol, vol. 59, no. 2, pp. 253-70, Jan 20 2014, doi:
10.1088/0031-9155/59/2/253.
[15] A. R. Smolock et al., "Robotically Assisted Sonic Therapy as a Noninvasive
Nonthermal Ablation Modality: Proof of Concept in a Porcine Liver Model,"
Radiology, vol. 287, no. 2, pp. 485-493, May 2018, doi:
10.1148/radiol.2018171544.
[16] N. R. Styn, J. C. Wheat, T. L. Hall, and W. W. Roberts, "Histotripsy of VX-2 tumor
implanted in a renal rabbit model," (in eng), J Endourol, vol. 24, no. 7, pp. 1145-
50, Jul 2010, doi: 10.1089/end.2010.0123.
[17] E. Vlaisavljevich et al., "Image-Guided Non-Invasive Ultrasound Liver Ablation
using Histotripsy: Feasibility Study in an In Vivo Porcine Model. ," Ultrasound
Med Biol, 2013.
[18] S. X. Vidal Jove J, Vlaisavljevich E, Cannata J, Duryea A, Miller R, Lee F and
Ziemlewicz T, "Phase I Study of Safety and Efficacy of Hepatic Histotripsy:
Preliminary Results of First in Man Experience with Robotically-Assisted Sonic
Therapy," in International Society of Therapeutic Ultrasound, Barcelona, Spain,
2019.
154
[19] S. Qu et al., "Non-thermal histotripsy tumor ablation promotes abscopal immune
responses that enhance cancer immunotherapy," Journal for ImmunoTherapy of
Cancer, vol. 8, no. 1, 2020.
[20] T. Worlikar et al., "Non-invasive liver cancer ablation using histotripsy in an in
vivo murine hepatocellular carcinoma (HCC) model," in Ultrasonics Symposium
(IUS), 2017 IEEE International, 2017: IEEE, pp. 1-1.
[21] T. F. Vandamme, "Use of rodents as models of human diseases," (in eng), J
Pharm Bioallied Sci, vol. 6, no. 1, pp. 2-9, 2014, doi: 10.4103/0975-7406.124301.
[22] J. J. Tentler et al., "Patient-derived tumour xenografts as models for oncology
drug development," (in eng), Nature reviews. Clinical oncology, vol. 9, no. 6, pp.
338-50, Apr 17 2012, doi: 10.1038/nrclinonc.2012.61.
[23] M. G. S. Consortium, "Initial sequencing and comparative analysis of the mouse
genome," Nature, vol. 420, no. 6915, p. 520, 2002.
[24] J. Rossant and C. McKerlie, "Mouse-based phenogenomics for modelling human
disease," Trends in molecular medicine, vol. 7, no. 11, pp. 502-507, 2001.
[25] K. Paigen, "A miracle enough: the power of mice," Nature medicine, vol. 1, no. 3,
p. 215, 1995.
[26] M. De Jong and T. Maina, "Of mice and humans: are they the same?—
Implications in cancer translational research," Journal of Nuclear Medicine, vol.
51, no. 4, pp. 501-504, 2010.
[27] T. Flisikowska, A. Kind, and A. Schnieke, "Pigs as models of human cancers,"
Theriogenology, vol. 86, no. 1, pp. 433-7, Jul 1 2016, doi:
10.1016/j.theriogenology.2016.04.058.
155
[28] K. M. Schachtschneider et al., "The Oncopig Cancer Model: An Innovative Large
Animal Translational Oncology Platform," Front Oncol, vol. 7, p. 190, 2017, doi:
10.3389/fonc.2017.00190.
[29] L. B. Schook et al., "A Genetic Porcine Model of Cancer," PLoS One, vol. 10, no.
7, p. e0128864, 2015, doi: 10.1371/journal.pone.0128864.
[30] K. Lee, K. Farrell, and K. Uh, "Application of genome-editing systems to enhance
available pig resources for agriculture and biomedicine," (in eng), Reprod Fertil
Dev, vol. 32, no. 2, pp. 40-49, Jan 2019, doi: 10.1071/rd19273.
[31] S. S. Pinho, S. Carvalho, J. Cabral, C. A. Reis, and F. Gartner, "Canine tumors: a
spontaneous animal model of human carcinogenesis," Transl Res, vol. 159, no.
3, pp. 165-72, Mar 2012, doi: 10.1016/j.trsl.2011.11.005.
[32] J. D. Schiffman and M. Breen, "Comparative oncology: what dogs and other
species can teach us about humans with cancer," Philos Trans R Soc Lond B
Biol Sci, vol. 370, no. 1673, Jul 19 2015, doi: 10.1098/rstb.2014.0231.
[33] M. Paoloni et al., "Defining the Pharmacodynamic Profile and Therapeutic Index
of NHS-IL12 Immunocytokine in Dogs with Malignant Melanoma," PLoS One, vol.
10, no. 6, p. e0129954, 2015, doi: 10.1371/journal.pone.0129954.
[34] M. Paoloni et al., "Prospective molecular profiling of canine cancers provides a
clinically relevant comparative model for evaluating personalized medicine
(PMed) trials," PLoS One, vol. 9, no. 3, p. e90028, 2014, doi:
10.1371/journal.pone.0090028.
[35] M. Paoloni, S. Lana, D. Thamm, C. Mazcko, and S. Withrow, "The creation of the
Comparative Oncology Trials Consortium Pharmacodynamic Core: Infrastructure
156
for a virtual laboratory," Vet J, vol. 185, no. 1, pp. 88-9, Jul 2010, doi:
10.1016/j.tvjl.2010.04.019.
[36] M. C. Paoloni et al., "Rapamycin pharmacokinetic and pharmacodynamic
relationships in osteosarcoma: a comparative oncology study in dogs," PLoS
One, vol. 5, no. 6, p. e11013, Jun 08 2010, doi: 10.1371/journal.pone.0011013.
[37] I. Gordon, M. Paoloni, C. Mazcko, and C. Khanna, "The Comparative Oncology
Trials Consortium: using spontaneously occurring cancers in dogs to inform the
cancer drug development pathway," PLoS Med, vol. 6, no. 10, p. e1000161, Oct
2009, doi: 10.1371/journal.pmed.1000161.
[38] M. C. Paoloni et al., "Launching a novel preclinical infrastructure: comparative
oncology trials consortium directed therapeutic targeting of TNFalpha to cancer
vasculature," PLoS One, vol. 4, no. 3, p. e4972, 2009, doi:
10.1371/journal.pone.0004972.
[39] C. A. London et al., "Preclinical evaluation of the novel, orally bioavailable
Selective Inhibitor of Nuclear Export (SINE) KPT-335 in spontaneous canine
cancer: results of a phase I study," PLoS One, vol. 9, no. 2, p. e87585, 2014, doi:
10.1371/journal.pone.0087585.
[40] J. E. Parsons, C. A. Cain, G. D. Abrams, and J. B. Fowlkes, "Pulsed cavitational
ultrasound therapy for controlled tissue homogenization," (in eng), Ultrasound in
Medicine and Biology, vol. 32, no. 1, pp. 115-29, Jan 2006, doi: S0301-
5629(05)00374-1 [pii] 10.1016/j.ultrasmedbio.2005.09.005.
[41] J. E. Parsons, C. A. Cain, and J. B. Fowlkes, "Cost-effective assembly of a basic
fiber-optic hydrophone for measurement of high-amplitude therapeutic ultrasound
157
fields," J Acoust Soc Am, vol. 119, no. 3, pp. 1432-40, Mar 2006, doi:
10.1121/1.2166708.
[42] E. Vlaisavljevich, T. Gerhardson, T. Hall, and Z. Xu, "Effects of f-number on the
histotripsy intrinsic threshold and cavitation bubble cloud behavior," Phys Med
Biol, vol. 62, no. 4, pp. 1269-1290, Feb 21 2017, doi: 10.1088/1361-
6560/aa54c7.
[43] H. R. Holmes et al., "Focused ultrasound extraction (FUSE) for the rapid
extraction of DNA from tissue matrices," (in English), Methods Ecol Evol, Oct 20
2020, doi: 10.1111/2041-210x.13505.
[44] J. Khirallah et al., "Nanoparticle-mediated histotripsy (NMH) using
perfluorohexane 'nanocones'," Phys Med Biol, vol. 64, no. 12, p. 125018, Jun 20
2019, doi: 10.1088/1361-6560/ab207e.
[45] R. M. Brock et al., "Patient derived xenografts expand human primary pancreatic
tumor tissue availability for ex vivo irreversible electroporation testing," Frontiers
in Oncology, vol. 10, 2020.
[46] D. N. Kwon et al., "Production of biallelic CMP-Neu5Ac hydroxylase knock-out
pigs," (in eng), Sci Rep, vol. 3, p. 1981, 2013, doi: 10.1038/srep01981.
[47] J. T. Kang et al., "Production of GGTA1 targeted pigs using TALEN-mediated
genome editing technology," (in Eng), Journal of veterinary science, Aug 4 2015.
[48] B. P. Beaton et al., "Inclusion of homologous DNA in nuclease-mediated gene
targeting facilitates a higher incidence of bi-allelically modified cells," (in eng),
Xenotransplantation, vol. 22, no. 5, pp. 379-90, Sep-Oct 2015, doi:
10.1111/xen.12194.
158
[49] K. Lee et al., "Engraftment of human iPS cells and allogeneic porcine cells into
pigs with inactivated RAG2 and accompanying severe combined
immunodeficiency," Proc Natl Acad Sci U S A, vol. 111, no. 20, pp. 7260-5, May
20 2014, doi: 10.1073/pnas.1406376111.
[50] S. Lei et al., "Increased and prolonged human norovirus infection in RAG2/IL2RG
deficient gnotobiotic pigs with severe combined immunodeficiency," Sci Rep, vol.
6, p. 25222, Apr 27 2016, doi: 10.1038/srep25222.
[51] Y. J. Choi et al., "Recombination activating gene-2(null) severe combined
immunodeficient pigs and mice engraft human induced pluripotent stem cells
differently," Oncotarget, vol. 8, no. 41, pp. 69398-69407, Sep 19 2017, doi:
10.18632/oncotarget.20626.
[52] J. T. Kang et al., "Generation of RUNX3 knockout pigs using CRISPR/Cas9-
mediated gene targeting," (in eng), Reproduction in domestic animals =
Zuchthygiene, vol. 51, no. 6, pp. 970-978, Dec 2016, doi: 10.1111/rda.12775.
[53] K. Lee et al., "Piglets produced from cloned blastocysts cultured in vitro with
GM‐CSF," Molecular reproduction and development, vol. 80, no. 2, pp. 145-154,
2013.
[54] L. Yuan, P. M. Jobst, and M. Weiss, "Gnotobiotic Pigs: From Establishing Facility
to Modeling Human Infectious Diseases," in Gnotobiotics: Elsevier, 2017, pp.
349-368.
[55] A. V. M. Association, "Animal Health Studies Database," ed, 2016.
[56] T. Worlikar et al., "Histotripsy for Non-Invasive Ablation of Hepatocellular
Carcinoma (HCC) Tumor in a Subcutaneous Xenograft Murine Model," in 2018
159
40th Annual International Conference of the IEEE Engineering in Medicine and
Biology Society (EMBC), 2018: IEEE, pp. 6064-6067.
[57] S. Lei et al., "Increased and prolonged human norovirus infection in RAG2/IL2RG
deficient gnotobiotic pigs with severe combined immunodeficiency," Scientific
reports, vol. 6, p. 25222, 2016.
[58] K. Lee et al., "Engraftment of human iPS cells and allogeneic porcine cells into
pigs with inactivated RAG2 and accompanying severe combined
immunodeficiency," Proceedings of the National Academy of Sciences, vol. 111,
no. 20, pp. 7260-7265, 2014.
[59] M. T. Basel et al., "Human xenografts are not rejected in a naturally occurring
immunodeficient porcine line: a human tumor model in pigs," BioResearch open
access, vol. 1, no. 2, pp. 63-68, 2012.
[60] A. K. LeBlanc et al., "Creation of an NCI comparative brain tumor consortium:
informing the translation of new knowledge from canine to human brain tumor
patients," Neuro Oncol, vol. 18, no. 9, pp. 1209-18, Sep 2016, doi:
10.1093/neuonc/now051.
[61] VT-VMCVM. "VT-VMCVM Website." http://www.vetmed.vt.edu/clinical-trials/
(accessed.
[62] M. L. Hauck and M. L. Oblak, "19 - Tumors of the Skin and Subcutaneous
Tissues," in Withrow and MacEwen's Small Animal Clinical Oncology (Sixth
Edition), D. M. Vail, D. H. Thamm, and J. M. Liptak Eds. St. Louis (MO): W.B.
Saunders, 2020, pp. 352-366.
160
[63] J. M. Liptak and N. I. Christensen, "22 - Soft Tissue Sarcomas," in Withrow and
MacEwen's Small Animal Clinical Oncology (Sixth Edition), D. M. Vail, D. H.
Thamm, and J. M. Liptak Eds. St. Louis (MO): W.B. Saunders, 2019, pp. 404-
431.
[64] J. C. Friebele, J. Peck, X. Pan, M. Abdel-Rasoul, and J. L. Mayerson,
"Osteosarcoma: A Meta-Analysis and Review of the Literature," Am J Orthop
(Belle Mead NJ), vol. 44, no. 12, pp. 547-53, Dec 2015. [Online]. Available:
https://www.ncbi.nlm.nih.gov/pubmed/26665241.
[65] J. M. Fenger, C. A. London, and W. C. Kisseberth, "Canine osteosarcoma: a
naturally occurring disease to inform pediatric oncology," ILAR J, vol. 55, no. 1,
pp. 69-85, 2014, doi: 10.1093/ilar/ilu009.
[66] S. J. Withrow, B. E. Powers, R. C. Straw, and R. M. Wilkins, "Comparative
aspects of osteosarcoma. Dog versus man," Clin Orthop Relat Res, no. 270, pp.
159-68, Sep 1991. [Online]. Available:
http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=
Citation&list_uids=1884536.
[67] M. Paoloni et al., "Canine tumor cross-species genomics uncovers targets linked
to osteosarcoma progression," BMC genomics, vol. 10, p. 625, 2009, doi:
10.1186/1471-2164-10-625.
[68] F. Mueller, B. Fuchs, and B. Kaser-Hotz, "Comparative biology of human and
canine osteosarcoma," Anticancer research, vol. 27, no. 1A, pp. 155-64, Jan-Feb
2007. [Online]. Available: http://www.ncbi.nlm.nih.gov/pubmed/17352227.
161
[69] E. Morello, M. Martano, and P. Buracco, "Biology, diagnosis and treatment of
canine appendicular osteosarcoma: similarities and differences with human
osteosarcoma," Vet J, vol. 189, no. 3, pp. 268-77, Sep 2011, doi:
10.1016/j.tvjl.2010.08.014.
[70] K. S. Rankin, M. Starkey, J. Lunec, C. H. Gerrand, S. Murphy, and S. Biswas, "Of
dogs and men: comparative biology as a tool for the discovery of novel
biomarkers and drug development targets in osteosarcoma," Pediatr Blood
Cancer, vol. 58, no. 3, pp. 327-33, Mar 2012, doi: 10.1002/pbc.23341.
[71] R. Gorlick and C. Khanna, "Osteosarcoma," J Bone Miner Res, vol. 25, no. 4, pp.
683-91, Apr 2010, doi: 10.1002/jbmr.77.
[72] J. L. Rowell, D. O. McCarthy, and C. E. Alvarez, "Dog models of naturally
occurring cancer," Trends Mol Med, vol. 17, no. 7, pp. 380-8, Jul 2011, doi:
10.1016/j.molmed.2011.02.004.
[73] L. Mirabello, R. J. Troisi, and S. A. Savage, "International osteosarcoma
incidence patterns in children and adolescents, middle ages and elderly
persons," Int J Cancer, vol. 125, no. 1, pp. 229-34, Jul 01 2009, doi:
10.1002/ijc.24320.
[74] S. F. Altekruse, K. A. McGlynn, and M. E. Reichman, "Hepatocellular carcinoma
incidence, mortality, and survival trends in the United States from 1975 to 2005,"
Journal of clinical oncology : official journal of the American Society of Clinical
Oncology, vol. 27, no. 9, pp. 1485-91, Mar 20 2009, doi:
10.1200/JCO.2008.20.7753.
162
[75] S. A. Shah, J. K. Smith, Y. Li, S. C. Ng, J. E. Carroll, and J. F. Tseng,
"Underutilization of therapy for hepatocellular carcinoma in the medicare
population," Cancer, vol. 117, no. 5, pp. 1019-26, Mar 01 2011, doi:
10.1002/cncr.25683.
[76] C. J. Sonnenday, J. B. Dimick, R. D. Schulick, and M. A. Choti, "Racial and
geographic disparities in the utilization of surgical therapy for hepatocellular
carcinoma," J Gastrointest Surg, vol. 11, no. 12, pp. 1636-46; discussion 1646,
Dec 2007, doi: 10.1007/s11605-007-0315-8.
[77] S. Chacko and S. Samanta, ""Hepatocellular carcinoma: A life-threatening
disease"," Biomed Pharmacother, vol. 84, pp. 1679-1688, Dec 2016, doi:
10.1016/j.biopha.2016.10.078.
[78] D. R. Strombeck, "Clinicopathologic features of primary and metastatic neoplastic
disease of the liver in dogs," J Am Vet Med Assoc, vol. 173, no. 3, pp. 267-9, Aug
01 1978. [Online]. Available: https://www.ncbi.nlm.nih.gov/pubmed/211107.
[79] J. M. Liptak et al., "Massive hepatocellular carcinoma in dogs: 48 cases (1992-
2002)," J Am Vet Med Assoc, vol. 225, no. 8, pp. 1225-30, Oct 15 2004. [Online].
Available: https://www.ncbi.nlm.nih.gov/pubmed/15521445.
[80] C. Weisse, C. A. Clifford, D. Holt, and J. A. Solomon, "Percutaneous arterial
embolization and chemoembolization for treatment of benign and malignant
tumors in three dogs and a goat," J Am Vet Med Assoc, vol. 221, no. 10, pp.
1430-6, 1419, Nov 15 2002. [Online]. Available:
https://www.ncbi.nlm.nih.gov/pubmed/12458612.
163
[81] Y. K. Cho, H. Rhim, and S. Noh, "Radiofrequency ablation versus surgical
resection as primary treatment of hepatocellular carcinoma meeting the Milan
criteria: a systematic review," J Gastroenterol Hepatol, vol. 26, no. 9, pp. 1354-
60, Sep 2011, doi: 10.1111/j.1440-1746.2011.06812.x.
[82] R. Z. Swan, D. Sindram, J. B. Martinie, and D. A. Iannitti, "Operative microwave
ablation for hepatocellular carcinoma: complications, recurrence, and long-term
outcomes," J Gastrointest Surg, vol. 17, no. 4, pp. 719-29, Apr 2013, doi:
10.1007/s11605-013-2164-y.
[83] M. Zhang, H. Ma, J. Zhang, L. He, X. Ye, and X. Li, "Comparison of microwave
ablation and hepatic resection for hepatocellular carcinoma: a meta-analysis,"
Onco Targets Ther, vol. 10, pp. 4829-4839, 2017, doi: 10.2147/OTT.S141968.
[84] N. Lucchina et al., "Current role of microwave ablation in the treatment of small
hepatocellular carcinomas," Ann Gastroenterol, vol. 29, no. 4, pp. 460-465, Oct-
Dec 2016, doi: 10.20524/aog.2016.0066.
[85] P. Liang and Y. Wang, "Microwave ablation of hepatocellular carcinoma,"
Oncology, vol. 72 Suppl 1, pp. 124-31, 2007, doi: 10.1159/000111718.
[86] D. J. Niemeyer et al., "Optimal ablation volumes are achieved at submaximal
power settings in a 2.45-GHz microwave ablation system," Surg Innov, vol. 22,
no. 1, pp. 41-5, Feb 2015, doi: 10.1177/1553350614532535.
[87] F. Wu et al., "Extracorporeal high intensity focused ultrasound ablation in the
treatment of 1038 patients with solid carcinomas in China: an overview," Ultrason
Sonochem, vol. 11, no. 3-4, pp. 149-54, May 2004, doi:
10.1016/j.ultsonch.2004.01.011.
164
[88] C. X. Li et al., "Analysis of clinical effect of high-intensity focused ultrasound on
liver cancer," World J Gastroenterol, vol. 10, no. 15, pp. 2201-4, Aug 1 2004, doi:
10.3748/wjg.v10.i15.2201.
[89] F. Wu et al., "Pathological changes in human malignant carcinoma treated with
high-intensity focused ultrasound," Ultrasound Med Biol, vol. 27, no. 8, pp. 1099-
106, Aug 2001, doi: 10.1016/s0301-5629(01)00389-1.
[90] B. R. Partridge et al., "High-Frequency Irreversible Electroporation for Treatment
of Primary Liver Cancer: A Proof-of-Principle Study in Canine Hepatocellular
Carcinoma," (in eng), J Vasc Interv Radiol, vol. 31, no. 3, pp. 482-491.e4, Mar
2020, doi: 10.1016/j.jvir.2019.10.015.
[91] N. Beitel-White, R. C. G. Martin, Y. Li, R. M. Brock, I. C. Allen, and R. V.
Davalos, "Real-time prediction of patient immune cell modulation during
irreversible electroporation therapy," (in eng), Sci Rep, vol. 9, no. 1, p. 17739,
Nov 28 2019, doi: 10.1038/s41598-019-53974-w.
[92] V. M. Ringel-Scaia et al., "High-frequency irreversible electroporation is an
effective tumor ablation strategy that induces immunologic cell death and
promotes systemic anti-tumor immunity," EBioMedicine, vol. 44, pp. 112-125, Jun
2019, doi: 10.1016/j.ebiom.2019.05.036.
[93] Z. M. Bamboat et al., "Human liver dendritic cells promote T cell
hyporesponsiveness," Journal of immunology, vol. 182, no. 4, pp. 1901-11, Feb
15 2009, doi: 10.4049/jimmunol.0803404.
165
[94] A. P. Jewell, "Is the liver an important site for the development of immune
tolerance to tumours?," Med Hypotheses, vol. 64, no. 4, pp. 751-4, 2005, doi:
10.1016/j.mehy.2004.10.002.
[95] P. Guha, J. Reha, and S. C. Katz, "Immunosuppression in liver tumors: opening
the portal to effective immunotherapy," Cancer Gene Ther, vol. 24, no. 3, pp.
114-120, Mar 2017, doi: 10.1038/cgt.2016.54.
[96] I. A. Siddiqui et al., "Induction of rapid, reproducible hepatic ablations using next-
generation, high frequency irreversible electroporation (H-FIRE) in vivo," HPB :
the official journal of the International Hepato Pancreato Biliary Association, vol.
18, no. 9, pp. 726-34, Sep 2016, doi: 10.1016/j.hpb.2016.06.015.
[97] C. M. Goutal, B. L. Brugmann, and K. A. Ryan, "Insulinoma in dogs: a review," J
Am Anim Hosp Assoc, vol. 48, no. 3, pp. 151-63, May-Jun 2012, doi:
10.5326/JAAHA-MS-5745.
[98] T. Schermerhorn, "Canine insulinoma as a model for studying molecular genetics
of tumorigenesis and metastasis," Vet J, vol. 197, no. 2, pp. 126-7, Aug 2013,
doi: 10.1016/j.tvjl.2013.04.017.
[99] M. Gillard et al., "Naturally occurring melanomas in dogs as models for non-UV
pathways of human melanomas," Pigment Cell Melanoma Res, vol. 27, no. 1, pp.
90-102, Jan 2014, doi: 10.1111/pcmr.12170.
[100] B. Hernandez, H. A. Adissu, B. R. Wei, H. T. Michael, G. Merlino, and R. M.
Simpson, "Naturally Occurring Canine Melanoma as a Predictive Comparative
Oncology Model for Human Mucosal and Other Triple Wild-Type Melanomas," Int
J Mol Sci, vol. 19, no. 2, Jan 30 2018, doi: 10.3390/ijms19020394.
166
[101] R. M. Simpson et al., "Sporadic naturally occurring melanoma in dogs as a
preclinical model for human melanoma," Pigment Cell Melanoma Res, vol. 27,
no. 1, pp. 37-47, Jan 2014, doi: 10.1111/pcmr.12185.
[102] M. M. Rahman et al., "Transcriptome analysis of dog oral melanoma and its
oncogenic analogy with human melanoma," Oncol Rep, vol. 43, no. 1, pp. 16-30,
Jan 2020, doi: 10.3892/or.2019.7391.
[103] A. Prouteau and C. Andre, "Canine Melanomas as Models for Human
Melanomas: Clinical, Histological, and Genetic Comparison," Genes (Basel), vol.
10, no. 7, Jun 30 2019, doi: 10.3390/genes10070501.
[104] W. Supsavhad, W. P. Dirksen, C. K. Martin, and T. J. Rosol, "Animal models of
head and neck squamous cell carcinoma," Vet J, vol. 210, pp. 7-16, Apr 2016,
doi: 10.1016/j.tvjl.2015.11.006.
[105] R. F. Hoyt and S. J. Withrow, "Oral malignancy in the dog.," Journal of the
American Hospital Association, vol. 20, pp. 83-92, 1984.
[106] R. J. Todoroff and R. S. Brodey, "Oral and pharyngeal neoplasia in the dog: a
retrospective survey of 361 cases," (in eng), J Am Vet Med Assoc, vol. 175, no.
6, pp. 567-71, Sep 15 1979. [Online]. Available:
http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=
Citation&list_uids=511751.
[107] J. K. Kosovsky, D. T. Matthiesen, S. M. Marretta, and A. K. Patnaik, "Results of
partial mandibulectomy for the treatment of oral tumors in 142 dogs," (in eng),
Vet Surg, vol. 20, no. 6, pp. 397-401, Nov-Dec 1991. [Online]. Available:
167
http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=
Citation&list_uids=1369522.
[108] N. Farcas, B. Arzi, and F. J. Verstraete, "Oral and maxillofacial osteosarcoma in
dogs: a review," Vet Comp Oncol, vol. 12, no. 3, pp. 169-80, Sep 2014, doi:
10.1111/j.1476-5829.2012.00352.x.
[109] J. Rossmeisl, "Maximizing Local Access to Therapeutic Deliveries in
Glioblastoma. Part V: Clinically Relevant Model for Testing New Therapeutic
Approaches," in Glioblastoma, S. De Vleeschouwer Ed. Brisbane (AU): Codon
Publications, 2017.
[110] M. F. Lorenzo et al., "Temporal Characterization of Blood-Brain Barrier Disruption
with High-Frequency Electroporation," (in eng), Cancers (Basel), vol. 11, no. 12,
Nov 23 2019, doi: 10.3390/cancers11121850.
[111] S. Demaria et al., "Ionizing radiation inhibition of distant untreated tumors
(abscopal effect) is immune mediated," (in eng), Int J Radiat Oncol Biol Phys, vol.
58, no. 3, pp. 862-70, Mar 1 2004, doi: 10.1016/j.ijrobp.2003.09.012.
[112] E. Vlaisavljevich, A. Maxwell, M. Warnez, E. Johnsen, C. Cain, and Z. Xu,
"Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
mechanical properties," (in eng), IEEE Transactions on Ultrasonics,
Ferroelectrics, and Frequency Control, vol. 61, no. 2, pp. 341-52, Feb 2014, doi:
10.1109/TUFFC.2014.6722618.
[113] E. Vlaisavljevich et al., "Effects of tissue stiffness, ultrasound frequency, and
pressure on histotripsy-induced cavitation bubble behavior," Phys Med Biol, vol.
60, no. 6, pp. 2271-92, Mar 21 2015, doi: 10.1088/0031-9155/60/6/2271.
168
[114] E. Vlaisavljevich, Z. Xu, A. Arvidson, L. Jin, W. Roberts, and C. Cain, "Effects of
thermal preconditioning on tissue susceptibility to histotripsy," Ultrasound in
medicine & biology, vol. 41, no. 11, pp. 2938-2954, 2015.
[115] H. Y. Ge et al., "Correlation between ultrasound reflection intensity and tumor
ablation ratio of late-stage pancreatic carcinoma in HIFU therapy: dynamic
observation on ultrasound reflection intensity," (in eng), ScientificWorldJournal,
vol. 2013, p. 852874, 2013, doi: 10.1155/2013/852874.
[116] H. Y. Ge et al., "High-intensity focused ultrasound treatment of late-stage
pancreatic body carcinoma: optimal tumor depth for safe ablation," Ultrasound
Med Biol, vol. 40, no. 5, pp. 947-55, May 2014, doi:
10.1016/j.ultrasmedbio.2013.11.020.
[117] K. E. Mitchell et al., "Outcomes of Limb-Sparing Surgery Using Two Generations
of Metal Endoprosthesis in 45 Dogs With Distal Radial Osteosarcoma. A
Veterinary Society of Surgical Oncology Retrospective Study," Vet Surg, vol. 45,
no. 1, pp. 36-43, Jan 2016, doi: 10.1111/vsu.12423.
[118] A. K. LeBlanc and C. N. Mazcko, "Improving human cancer therapy through the
evaluation of pet dogs," (in eng), Nat Rev Cancer, vol. 20, no. 12, pp. 727-742,
Dec 2020, doi: 10.1038/s41568-020-0297-3.
169
Chapter 5. Establishing a SCID-Like Porcine Model of Human Cancer for Novel Therapy Development with Pancreatic Adenocarcinoma and Irreversible
Electroporation
Alissa Hendricks-Wenger, Kenneth N. Aycock, Margaret A. Nagai-Singer, Sheryl Coutermarsh-Ott, Melvin F. Lorenzo, Jessica Gannon, Kyungjun Uh, Kayla Farrell,
Natalie Beitel-White, Rebecca M. Brock, Alexander Simon, Holly A. Morrison, Joanne Tuohy, Sherrie Clark-Deener, Eli Vlaisavljevich, Rafael V. Davalos, Kiho Lee, Irving C.
Allen
This chapter has been previously published and used here with permission from: Hendricks-Wenger et al., Establishing a SCID-Like Porcine Model of Human Cancer for
Novel Therapy Development with Pancreatic Adenocarcinoma and Irreversible Electroporation, Scientific Reports, 2020, Nature Research
170
5.1 Abstract
New therapies to treat pancreatic cancer are direly needed. However, efficacious
interventions lack a strong preclinical model that can recapitulate patients’ anatomy and
physiology. Likewise, the availability of human primary malignant tissue for ex vivo
studies is limited. These are significant limitations in the biomedical device field. We
have developed RAG2/IL2RG deficient pigs using CRISPR/Cas9 as a novel, large
animal model for use in cancer xenograft studies of human pancreatic adenocarcinoma.
In this proof-of-concept study, these novel pigs were successfully generated using on-
demand genetic modifications in embryos, circumventing the need for breeding and
husbandry. Human Panc01 cells injected subcutaneously into the ears of RAG2/IL2RG
deficient pigs demonstrated 100% engraftment with growth rates similar to those
typically observed in mouse models. Histopathology revealed no immune cell infiltration
and tumor morphology was highly consistent with the mouse models. The electrical
properties and response to irreversible electroporation of the tumor tissue were found to
be similar to excised human pancreatic cancer tumors. The ample tumor tissue
produced enabled improved accuracy and modeling of the electrical properties of tumor
tissue. Together, this suggests that this model will be useful and capable of bridging the
gap of translating therapies from the bench to clinical application.
171
5.2 Keywords
Pancreatic Cancer; CRISPR/Cas9; Pig; Tumor Ablation; IRE
172
5.3 Introduction
Despite relatively low rates of occurrence, pancreatic cancer accounts for a
disproportionately high rate of cancer associated death. Surgery may cure patients with
Stage I disease; however, more than 75% of patients will present with borderline or
unresectable disease (Stages II-IV).1,2 Chemotherapy and radiation treatment protocols
result in limited responses and are rarely curative.3-6 Thus, new therapies are direly
needed. Over the last decade, tumor ablation strategies, such as radiofrequency,
microwave, and cryoablation have emerged as promising therapeutic options for
pancreatic cancer.7-9 Prior to human trials, the ablation modalities developed to date for
pancreatic cancer were originally developed using models based on data generated
from healthy tissues, rodent models, and occasionally spontaneous tumors in veterinary
patients. Unfortunately, there are major limitations with each of these approaches that
have hindered tumor ablation device development and limited data necessary for
accurate treatment planning in human patients. Thus, the lack of clinically and
physiologically relevant pre-clinical pancreatic cancer models continues to be a
significant limitation.
Murine models are the dominant in vivo model in the biomedical engineering and
cancer biology fields and are essential in acquiring fundamental data. Mice are easy to
house and handle, relatively inexpensive, share many genetic similarities to humans,
and there are many transgenic and knock-out lines available to study specific disease
mechanisms, proteins, and pathways.10-15 While these advantages make murine models
173
appealing, the gap between mice and humans is wide enough to prevent many of the
devices and therapies developed solely in mouse models from being safe and effective
in human patients. There are important anatomical differences between humans and
mice. Humans are approximately 2,500 times larger than mice, which accounts for
differences in many biological aspects, including metabolic rates, heart rates, and life
expectancy.15-19 Looking at carcinogenesis, mice develop malignant tumors more
readily than humans, due to significant differences in DNA repair and telomerase
regulation, among others.20 In an attempt to bridge this gap, more researchers are using
patient-derived tumor xenograft (PDX) models for oncological studies. In these models,
human tumors are engrafted into immunocompromised mice, allowing the local
microenvironment of the human tumor to be recapitulated in the mouse.11,21-24 While the
PDX model is an impressive improvement to mouse models, the anatomical differences
between mice and humans still presents a barrier to clinical relevance.
Porcine models have demonstrated increasing clinical relevance due to their similar
size, anatomy, and physiology to humans. Their large size is ideal and practical for
imaging technology studies and clinical device development actical.25 Porcine models
have been established as clinically-relevant models for toxicology, surgical procedures,
and designing equipment and instruments.26-28 Anatomical differences are especially
critical for the development and testing of medical devices for surgical and ablation
procedures. For instance, the safety and efficacy of minimal and non-invasive ablation
techniques such as radiofrequency ablation, microwave ablation, cryoablation,
irreversible electroporation (IRE), and focused ultrasound (HIFU, histotripsy) are difficult
to test in mice due to these anatomical differences. As a result, these devices are often
174
tested in healthy large animal models, which can provide important data on treatment
safety and energy delivery, but make it difficult to assess the effects of the therapies in
diseased tissues. Therefore, the development of large animal tumor models would be
beneficial for pre-clinical safety and efficacy studies. Overall, pigs as a model are good
at recapitulating human cancers due to their similar size, genome, metabolism,
telomerase activity, and organ micro- and macro-structures.29,30 However, unlike murine
models, existing porcine models are not well-established or reproducible from subject-
to-subject.
The availability of abundant, high quality tumor tissue is a major limitation in the
development of medical devices that require ex vivo modeling and validation. This is
especially true in pancreatic cancer research, where human specimens from patients
are typically highly limited and restricted to small biopsies. Likewise, while post-mortem
tissue is more available, the quality of the tissue is often not ideal. While mice are
excellent bio-incubators for human tissue and cell lines, the size of tumor tissue is often
limited due to the relatively small size of the animal. We have found this to also be a
significant limitation in the development of irreversible electroporation (IRE) based
therapeutic strategies. While healthy porcine tissues and mouse models have been
critical for therapeutic development, we have found the lack of appropriate pancreatic
cancer tissues to be a limitation. It has long been known that electrical conductivity
deviates from its physiological value once tissues are removed from the body, but most
tissues remain minimally changed within 1 hour of removal.31
Members of our research team co-invented IRE as a non-thermal tissue ablation
modality capable of treating solid tumors32. IRE involves delivering a series of low
175
energy, unipolar electric pulses through electrodes inserted directly into the tumor. The
induced electric field distribution produces structural defects in the target cell membrane
that cause cell death. IRE has been successfully adapted for the treatment of tumors
considered, until now, inoperable due to their proximity to critical structures such as
blood vessels, nerves, and ducts.33-36
In an effort to circumvent the limitations related to animal models of pancreatic
cancer and further advance the optimization of IRE utilizing human pancreatic cancer
tissue, we developed an immunodeficient porcine model that has proven ideal for
human cell transplantation and xenograft studies.37-41 Utilizing CRISPR/Cas9, we have
previously generated pigs with targeted deletions of RAG2/IL2RG, and established that
these pigs are immunodeficient and thrive under our unique gnotobiotic housing
conditions.41 This prior study utilized these pigs to study the pathogenesis of a human
virus.41 Expanding upon these previous studies, here we deploy the RAG2/IL2RG
deficient pigs as a novel tool to study pancreatic cancer. For the present study, we
describe the original proof-of-concept studies, utilizing these pigs to propagate the
human Panc01 pancreatic cancer cell line for use in ex vivo IRE modeling. This strategy
generated an ample supply of high-quality pancreatic tumor tissue for electrical property
modeling and IRE assessments. We believe that this model is a promising step towards
studying human cancer treatments in a highly-relevant system with the potential for
significant clinical impact.
176
5.4 Materials And Methods
5.4.1 Generation of Immunodeficient Pigs
Table 5-1 Target sequences of RAG2 and IL2RG sgRNAs.
Target gene Sequence (PAM)
RAG2 5’ - GAATGACCATATCTGCCTTC(AGG)
5’ - GTATAGTCGAGGGAAAAGTA(TGG)
IL2RG
5’ - CCAACCTAACTCTGCACTAC(TGG)
5’ - TGGAAACGTTGAGAGTCCC(AGG)
5’ - AAACGTTGAGAGTCCCAGG(GGG)
Oocytes aspirated from sow ovaries were matured in maturation medium (TCM-199
medium supplemented with 0.5 IU ml−1 FSH, 0.5 IU ml−1 LH, 0.82 mM cysteine, 3.02 mM
glucose, 0.91 mM sodium pyruvate and 10 ng ml−1 EGF) for 40 – 44 hours. After in vitro
maturation, cumulus cells were removed by vortexing in the presence of hyaluronidase,
then mature oocytes extruding the first polar body were transferred to fertilization media
(modified Tris-buffered medium supplemented with 113.1 mM NaCl, 3 mM KCl, 7.5 mM
CaCl2, 11 mM glucose, 20 mM Tris, 2 mM caffeine, 5 mM sodium pyruvate and
2 mg ml−1 BSA). Extended semen was washed with PBS, then introduced into the IVF
dishes containing oocytes. The gametes were co-incubated for 5 hours at 38.5 °C and
5% CO2. Microinjection was conducted in manipulation media (TCM199 with 0.6 mM
NaHCO3, 2.9 mM HEPES, 30 mM NaCl, 10 ng ml−1 gentamicin and 3 mg ml−1 BSA)
covered with mineral oil on the heated stage of a Nikon inverted microscope (Nikon
177
Corporation, Tokyo, Japan). Two sgRNAs targeting the RAG2 gene, three sgRNAs
targeting the IL2RG gene, and Cas9 mRNA (10 ng/µl sgRNA each and 20 ng/μl Cas9
mRNA) were introduced into the cytoplasm of the presumable zygotes using a FemtoJet
microinjector (Eppendorf, Hamburg, Germany). Target sequences of the sgRNAs can
be found in Table 1. The injected embryos were moved to culture media supplemented
with 10 ng ml−1 GM-CSF and incubated at 38.5 °C, 5% CO2, and 5% O2 until embryo
transfer 57. The cultured embryos were transferred into a surrogate gilt at day 4 post-IVF
by surgically transferring into the oviduct. Pregnancy was determined by ultrasound at
around day 30 of gestation. To genotype the mutations generated by CRISPR/Cas9
system, genomic DNA was isolated from tail-tips of the newborn piglets using PureLink
Genomic DNA kit (Thermo Fisher Scientific) following the manufacturer’s instructions.
Primers flanking projected double strand break (DSB) sites were designed to amplify
target region of RAG2 and IL2RG genes (Table 2). The target regions were PCR
amplified using Dream Taq DNA Polymerase (Thermo Fisher Scientific). PCR
conditions were as follows: initial denature at 95 °C for 2 min; denature at 95 °C for
30 sec, annealing at 60 °C for 30 sec, and extension at 72 °C for 30 sec for 34 cycles;
72 °C for 5 min; and holding at 4 °C. The PCR amplicons were sequenced to determine
the mutation types generated by the CRISPR/Cas9 system. Immunocompromised state
was verified with flow cytometry to analyze the presence B (CD79A), T (CD3E), and NK
(CD3E-/CD56+) cells.
178
Table 5-2 Primers used to amplify sgRNA target regions.
Primer Sequence
RAG2_Forward GGCTTTCCCATAACCTGGATATTGGGTC
RAG2 Reverse CGTCTCAGACTCATCTTCCTCATCATCTT
IL2RG_Forward CGGTAATAATCATGACTAGAGGGAATGAAAGATTGATTTATC
IL2RG_Reverse GAGAGAAAGTTGGGTGTCTATAAAAGAAGGGAGAATTAAAACTG
5.4.2 Animal Care
Six pigs (ntotal=5 males and ntotal=1 female; n=2 male and n=1 female used for the
current study) were delivered through a sterile hysterectomy process previously
described by a board-certified large animal veterinarian.58 Pigs were monitored regularly
until fully recovered from anesthesia and able to eat independently. Tail snips were
taken from each piglet while still under the influence of anesthesia for genotyping, and
ears were clipped for numbering. The pigs were born into sterile, gnotobiotic isolators
and were fed sterile boxed milk throughout the study in order to prevent infections.58 All
animal experiments were approved and carried out in accordance with the Virginia Tech
Institutional Animal Care and Use Committee under IACUC protocol: 19-117-CVM.
5.4.3 Tumor Injection and Monitoring
Human Panc01 cells (National Cancer Institute DTP, DCTP Tumor Repository)
were grown up in DMEM supplemented with 10% FBS and 1% penicillin/ streptomycin
and removed from plates with Trypsin in EDTA. While the pigs were still under the effect
179
of anesthesia from the hysterectomy, 1.2x106 Panc01 cells in 100 µL of Matrigel
(Corning, USA) were injected subcutaneously behind the ears of three pigs, with an
additional 100 µL of Matrigel behind the left ear. Tumors were measured three times per
week with plastic Vernier calipers. Diameter was calculated as the square root of the
product of the widest diameter and the diameter perpendicular to that. At the same point
of tumor measurement, weight and other health monitoring was also completed.
Following euthanasia, animals were necropsied and tissues were fixed in 10% formalin
for at least 24 hours. Paraffin-embedded formalin-fixed tissues were H&E stained and
evaluated by board-certified veterinary pathologist.
5.4.4 Immunohistochemistry
Pan02, PDX, and Panc01 samples (n=3/group) were subject to IHC-P staining of
CK19 (Thermo Fisher, MA5-12663). Paraffin-embedded formalin-fixed tissue sections
were deparaffinized following established xylene:ethonal wash protocol.59 Antigen
retrieval was achieved through 1-hour incubation at 95oC on a hot plate in sodium
citrate buffer with pH6 based upon the antibody manufacture’s suggestion. Once
cooled, slides were washed twice in tris-buffered saline (TBS) plus 0.025% Triton X-100
with gentle agitation. Primary antibody was diluted (1:250) in TBS with 1% BSA and
10% FBS overnight at 4 oC. Slides were washed four times in tris-buffered saline (TBS)
plus 0.025% Triton X-100 with gentle agitation, prior to 2 hours room temperature
incubation with secondary antibody (Cell Signaling, 7076P2) diluted (1:2000) in TBS
with 1% BSA and 10% FBS. Slides were washed three times for 5 minutes in TBS, then
180
developed with DAB substrate (Abcam, ab64238) following manufactures guidelines
and counterstained with eosin and methyl blue.
5.4.5 Ultrasound Imaging
Animals were anesthetized with Xylazine/Telazol. Prior to imaging, pigs were
cleaned and dried, and had the area of interest shaved to remove hair that could
interfere with imaging. Ultrasound imaging was done using both linear and phased array
ultrasound transducers (L18-10L30H-4 and P8-3L0S1-6, TELEMED, Lithuania, EU).
Images were acquired on a laptop computer and saved for future analysis. Targets were
confirmed by a board-certified veterinarian.
5.4.6 Tissue Handling and Electroporation
Small rectangular sections of healthy or malignant tissue were excised, washed to
prevent blood coagulation, and placed in modified PBS to help maintain osmotic
pressure balance as previously described.60 Immediately prior to pulsing, a sample of
the tissue was removed from PBS, sectioned into smaller pieces, and placed into an
insulating cylindrical polydimethylsiloxane (PDMS) mold as previously described.61 The
mold had radius of 3 mm and thickness of 0.56 cm. The tissue-containing mold was
placed between a set of parallel plate electrodes connected to a pulse generator (BTX
ECM 830, Harvard Apparatus, Holliston, MA). Baseline electrical conductivity was
measured by delivering a low voltage (~20 V) potential across the electrodes. Electrical
impedance was then recorded from 1 to 106 Hz.
181
All excised tumor and healthy tissues were analyzed within 1 hour of resection. Prior
to data analysis, all DC offsets were removed and a Butterworth filter was applied to
smooth the data and allow for automated analysis in MATLAB (MathWorks Inc., Natick,
MA, USA). To assess differences in electrical properties between our RAG2/IL2RG pig
Panc01 tumor model and primary human pancreatic cancer specimen, we compared
the initial tissue conductivities and the changes in tissue conductivity at 2.5 kV/cm.
Baseline electrical properties were recorded for healthy pancreas, prostate and brain
tissue excised from RAG2/IL2RG pigs. Prior to IRE delivery, a low voltage (~20 V) pulse
was administered to record sample-specific initial conductivity. This low potential pulse
does not cause electroporation-induced changes in electrical properties, and is thus
only diagnostic. The resulting value was used as a baseline to determine percent
differences in tissue conductivity during the subsequent high voltage, 100-pulse IRE
protocol. An electric field strength of either 500, 750, 1000, 1500, 2000, 2500, and 3,000
V/cm was delivered to each sample (n=1 for each electric field strength, n=7 total
samples). For comparison, the data acquired in this study are reported alongside tissue
conductivity data from primary human pancreatic cancer, as previously reported in
Beitel-White et al., primary pancreatic ductal adenocarcinoma (PDAC) tissue
propagated in mice from Brock et al., as well as, data obtained from literature for
healthy pancreas, prostate, and brain tissue.24,46,62-68
To monitor changes in temperature due to joule heating, a general-purpose STB
fiber optic thermometry probe (LumaSense, Santa Clara, CA, USA) was inserted at the
midpoint of the cylindrical tissue sample. Temperature was recorded at a frequency of 2
Hz using a Luxtron m3300 Biomedical Lab Kit (LumaSense, Santa Clara, CA, USA). An
182
unpaired t-test with Welch’s correction was used to compare the mean initial
conductivity.
Figure 5-1 Generation of RAG/ IL2RG knockout pigs. (A) Schematic approach for RAG2/IL2RG knockout pig production. First, the sgRNA and Cas9 mRNA was synthesized via in vitro transcription. Second, an optimal concentration of sgRNA and Cas9 was injected into presumable zygotes. Third, the injected embryos were transferred into a surrogate gilt. At the end of gestation, piglets were born via hysterectomy and maintained in gnotobiotic isolators. (B) Summary of piglet genotypes in IL2RG and RAG2 gene. Six piglets were born and analyzed. During PCR analysis, RAG2 was not amplified from the Piglet #2 DNA sample, suggesting a large deletion. Biallelic genotype indicates two differently modified alleles. Homozygous indicates one type of modified alleles. (C) Genotype of Piglet 1 as a representation of small deletions or insertions introduced by the CRISPR/Cas9 system.
183
5.5 Results
5.5.1 Generation of immunodeficient pigs carrying targeted deletions in RAG2/IL2RG
using the CRISPR/Cas9 system
Previous studies have utilized immunodeficient pigs in infectious disease studies
and to evaluate engraftment of human iPS cells.42-44 However, these pigs have yet to be
widely utilized in cancer research models. CRISPR/Cas9 RNAs were introduced into the
cytoplasm of in vitro fertilized pig oocytes (Fig. 1A). To disrupt pig immune system, the
RAG2 and IL2RG genes were targeted.42 In vitro fertilized oocytes (n = 454) were
injected with the targeted RNA for the CRISPR/Cas9 system, then cultured in vitro. On
day 4 post-IVF, 155 embryos at between the 8-cell and morula stage were transferred
to a surrogate gilt and subsequently six piglets were derived by hysterectomy. An
important advantage of this strategy is the ability to generate the RAG2/IL2RG deficient
pigs “on-demand”, without the need to maintain breeding colonies. Once born, the
immunocompromised state of the pigs requires them to be maintained in gnotobiotic
isolators under germ-free condition. Genotyping was conducted using genomic DNA
collected from tail-snips. Similar to our previous report,41 genotyping revealed that none
of the six piglets have wild-type sequences in either of the RAG2 or IL2RG genes (Fig.
1B detailed in Supplementary Fig. S1). All six piglets carried small insertion or deletion
mutations in both RAG2 and IL2RG genes (Fig. 1B). The genotype of piglet 1 is shown
as an example (Fig. 1C). A RAG2 fragment flanking the CRISPR/Cas9-induced DSBs
site could not be amplified from one piglet by PCR, suggesting a larger deletion caused
184
by CRISPR/Cas9 system. For tumor engraftment studies, 3 of the 6 pigs were randomly
selected for subsequent Panc01 studies. Aligned with the RAG2/IL2RG knockout
establishing study,42 the piglets presented B-T-NK-SCID phenotype (Fig. 2).
185
Figure 5-2 Confirmation of SCID-Like Phenotype. FACS analysis to detect the presence of lymphocytes in RAG2/IL2RG knockout piglets. Compared to wild-type control, SCID piglet possessed lower level of B (CD79A), T (CD3E), and NK (CD3E-/CD56+) cells.
5.5.2 Panc01 tumors successfully engrafted in all pigs carrying the targeted
RAG2/IL2RG deletion
Figure 5-3 Successful Engraftment of Panc01 Cells. (A) Tumor growth was measured three times per week for each pig. All animals developed palpable tumors that continued to grow over the course of the study. (B) Pig weight was monitored throughout the duration of the study and utilized as a surrogate for animal morbidity.
The purpose of the current study was to generate human tumors of sufficient size
and quality in our unique immunocompromised pigs for ex vivo assessments of IRE.
However, due to this being the first use of these pigs to propagate tumor tissue and the
potential for incomplete knockout of the immune system due to inefficiencies in the
CRISPR/Cas9 strategy, we first established engraftment proficiency and growth rates
for the Panc01 xenografts. Within 24 hours of the initial injection, tumor growth was
186
observed for all Panc01 tumors. After an initial period of rapid growth, within 5 days of
xenograft, the rate of increase stabilized and steadily increased to the target diameter of
1.0 - 1.6cm by day 36 (Fig. 3A). No changes were observed in morbidity, mortality, or
clinical parameters associated with tumor progression demonstrated here by the
change in pig weight over time (Fig. 3B). All tumors were easily palpated and easily
observed by day 36 (Fig. 4A). The Matrigel control injection was not detected (Fig. 4B).
Additional assessments utilizing ultrasound imaging revealed clearly defined tumors,
with clear delineations from surrounding tissues (Fig. 4C). Necropsy revealed the
subcutaneous tumors remained within the dermal layer, and where easily palpated and
observable from underneath the dermal layers (Fig. 4D). Tumor measurements
collected extra-dermally, sub-dermally, and with ultrasound were comparable for each
tumor (Fig. 4A, C-E). At gross necropsy, no evidence of metastasis was identified in
draining or other lymph nodes, nor in organs with major vascular supply, such as the
spleen, liver, or lung.
187
Figure 5-4 Subcutaneous Tumors Generated Ample, High-Quality Pancreatic Cancer Tissue for Subsequent Ex Vivo Tumor Ablation Assessments. (A) Tumor growth was readily observable above the skin behind the ear where the Panc01 cells were injected. (B) No observable foci were located in regions where Matrigel alone (control) was injected. (C) Ultrasound image confirmed the volumetric growth of the tumors. The tumor is outlined here in white for clarity. (D-E) Necropsy of the tumors confirmed that (D) they were confined to the dermis and (E) approximately the same size as measured extra-dermally.
5.5.3 Panc01 tumors are histopathologically similar to Pan02 and PDX tumors
generated in mice
Previous studies have utilized human xenografts in immunocompromised mice or
murine tumor injection into immunocompetent animals to generate tissue for ex vivo IRE
studies. However, the small size of the mice is a restriction for tumor growth and
limitation for any ultimate orthotopic studies. Due to the historic use of mice in IRE and
other tumor ablation modality development, we next compared the Panc01 tumors from
the RAG2/IL2RG pigs with Pan02 tumors generated in immunocompetent mice. In
188
Figure 5-5 Histopathology Comparison of Pre-Clinical Pancreatic Cancer Animal Models. Human PDX, mouse Pan02, and human Panc01 were compared. (A) Subcutaneous tumors in the mouse Pan02 model are densely cellular often with central areas of necrosis (asterisk). Individual cells are spindle-shaped and arranged in vague, irregular streams. (B) Subcutaneous tumors in the PDX mice are also highly cellular and also exhibit foci of necrosis (asterisk). At higher magnification, these cells are more polygonal and are often arranged to form irregular tubular structures with lumens containing inflammatory cells and debris (arrows). (C) Subcutaneous tumors in the porcine Panc01 model are also densely cellular and variably exhibit foci of necrosis. Individual cells are more polygonal in shape and rarely attempt to recapitulate glandular structures (arrows).
189
Figure 5-6 Immunohistochemical Confirmation of Ductal Cells in Pre-Clinical Pancreatic Cancer Animal Models. (A) Human PDX, (B) mouse Pan02, and (C) human Panc01 were compared for CK19 expression. Brown color indicates positive color. Nuclei are stained in dark blue, and remaining cell features are highlighted in light blue.
190
general, cell line tumors from both animal models were highly similar. However, as
expected simply due to the immune system status of each animal, we did observe some
minor differences in tumor histopathology. In the murine Pan02 and in the murine PDX
subcutaneous pancreatic adenocarcinoma models, nodular tumors develop at the site
of injection (Fig. 5A and 5B, respectively). The Pan02 tumors are densely cellular with
minimal fibrovascular stroma and often variably-sized foci of necrosis develop centrally
within the tumor. Tumor cells are spindle in shape and tend to be arranged in vague,
indistinct streams. Similar to most malignant tumor cells of any origin, they often exhibit
marked pleomorphism (differences in cell and nuclear size and shape across the
population) with numerous mitotic figures scattered throughout. In the PDX model, cells
derived from human tumors are injected into immunodeficient NOD-Scid gamma (NSG)
mice. Microscopically, the tumors in these mice look different from those in the Pan02
model. For example, in the PDX model, tumor cells are more polygonal in shape
(characteristic of epithelial cells) and are often arranged into disorganized glandular
structures. In the RAG2/IL2RG pig xenografts, microscopically the tumors are
characterized by dense cellularity on minimal fibrovascular stroma, as seen in other
models (Fig. 5C). Tumor cells, resemble an intermediate between the two murine
models. Cells are more polygonal in shape and do rarely attempt to recapitulate
glandular structures. These are often indistinct and do not often exhibit lumens.
Localization of CK 19, an established ductal cell marker, was also analyzed in Pan02,
PDX, and Pan01 tumors in order to establish the ductal characteristics of the tissues.45
Murine Pan02 sections were not found to express CK19 (Fig. 6A). In the murine PDX
191
and RAG2/IL2RG pig xenografts, CK19 expression was found throughout the tissues,
localizing in glandular and glandular-like structures respectively (Fig. 6B-C).
5.5.4 Panc01 tumors demonstrate similar electrical properties to primary patient derived
pancreatic cancer tissue and healthy porcine tissues
Figure 5-7 Primary human, SCID-like Panc01, and murine PDX tumors demonstrate similar conductivity increases following high voltage pulsed electric field. Ex vivo tissue samples directly from patients, RAG2/IL2RG animals, or NSG mice were sectioned and fit within a PDMS mold to impose a cylindrical shape factor. After sample preparation, individual samples were exposed to IRE pulses with varying electric field magnitudes. A) Summary of initial conductivities calculated from normal and cancerous SCID-like (black), human (red), and murine PDX (gray) tissue. B) The conductivity for each tissue type increased with the applied electric field. C) Percent increase in tissue conductivity at varying fields was determined from the sample-specific pre-pulse values. D) The adjusted conductivity is calculated from the percent difference values; here, the conductivities are normalized to the average initial tissue conductivity and adjusted based on the percent change.
192
Figure 5-8 Impedance for healthy tissue excised from the RAG2/IL2RG pigs correlate adult swine and humans. A Gamry Reference 600 potentiostat (Gamry, Warminster, PA, US) was used to record tissue impedance within a frequency range of 103 to 106 Hz. The real part of the tissue impedance was used in the calculation of tissue conductivity for further comparison to values previously reported in literature. A) Comparison of electrical conductivity from SCID-like pig tissue to healthy porcine (pancreas/brain) or human (prostate) tissue. The B) pancreatic, C) brain, and D) prostate tissue demonstrate similar electrical behavior to their normal counterparts across a wide frequency range. Each data point (dot) represents reported values for normal tissue conductivity, while the black line represents data gathered in this study and the red line represents normal porcine tissue. (*p < 0.05).
There are several significant advantages of the Panc01 porcine xenograft model
over other currently existing models and even primary human pancreatic cancer tissue.
Specifically, the Panc01 tumors are readily available, high quality, and in sufficient
193
volume for robust ex vivo studies. Thus, we next compared the electrical properties of
the Panc01 tissue with primary human pancreatic cancer tissue (PHPC) and murine-
propagated primary pancreatic ductal adenocarcinoma (PDAC) tissue as originally
reported in Brock et al. (Fig 7).24 The PHPC tissue (n = 7), excised Panc01 tumor
tissue (n = 7), and murine PDX tissue (n = 9) exhibited initial conductivities of 0.231
0.058 S/m, 0.242 0.043 S/m, and 0.225 0.070 S/m, respectively (Fig. 7A). No
statistically significant differences were found between these values. With an applied
electric field of 2.5 kV/cm, the mean intra-pulse conductivity increased to 0.288 S/m,
0.331 S/m, and 0.386 S/m for the PHPC, Panc01, and murine PDX tissue, respectively
(Fig. 7B). The relationship between the treated tumors is more clearly seen as a percent
increase from base line, with corresponding adjusted conductivity (Fig. 7C-D).
Directly comparing the healthy porcine pancreas in RAG2/IL2RG pigs to human
pancreas, the initial conductivity was 0.133 0.0059 S/m and 0.197 0.103 S/m (p =
0.0678), respectively (Fig. 7A). In addition to the healthy pig pancreas, we also
compared results with healthy brain and prostate. The EIS results for healthy brain,
pancreas, and prostate are reported as low frequency (10 kHz) and high frequency (1
MHz) measurements for direct comparison to literature values; the impedance spectrum
for each tissue and the comparison to literature values are shown in Figure 8. At low
frequency, the conductivity values for healthy tissue analyzed from the RAG2/IL2RG
model for brain (n = 8), pancreas (n = 3), and prostate (n = 7) tissue measured at 0.175
0.017 S/m, 0.145 0.005 S/m, and 0.343 0.039 S/m. In this same frequency, the
conductivity data measured from the standard healthy porcine model for brain (n = 5)
and pancreatic (n = 4) tissues measured at 0.167 0.016 S/m and 0.188 0.02 S/m,
194
respectively (Fig. 8A). A t-test was used to determine statistically significant differences
between the RAG2/IL2RG pig healthy tissue and the standard porcine model healthy
tissue; no statistical differences were detected for brain tissue (p = 0.4401), but a
statistical difference was detected for the pancreatic tissue (p = 0.021). High frequency
conductivity values for healthy tissue analyzed from the RAG2/IL2RG pig model for
brain (n = 8), pancreas (n = 3), and prostate (n = 7) tissue measured 0.294 0.021 S/m,
0.371 0.009 S/m, and 0.508 0.043 S/m (Fig. 8A). In this same frequency, conductivity
of the standard healthy porcine model for brain (n = 5) and pancreatic (n = 4) tissue
measured at 0.276 0.016 S/m and 0.383 0.009 S/m, respectively. Due to the lack of
easily accessible, normal porcine prostatic tissue, we compared SCID-like prostate
conductivity to that collected from freshly excised human prostates and reported in
Halter et. al, 2007.46 Halter and colleagues measured low- and high-frequency
conductivities of 0.43 0.27 S/m and 0.60 0.32 S/m, respectively, neither of which are
significantly different (p = 0.514 and p = 0.551) than the values reported herein for
SCID-like prostate conductivity (Fig. 8A). A t-test was used to determine statistically
significant differences between the RAG2/IL2RG healthy tissue and the standard
porcine model healthy tissue; no statistical differences were detected for brain (p =
0.1092) or pancreatic tissue (p = 0.1349) across porcine models. Due to the lack of
healthy porcine prostate data available, no direct comparisons were made between the
SCID-like prostate and conventional porcine prostate tissue at the abovementioned
frequencies.
195
5.6 Discussion
With their larger size compared to rodents and controlled immune system
knockdown, RAG2/IL2RG pigs created with the CRISPR/Cas9 system may be used as
a novel oncology model for translating developing therapies. They offer the opportunity
to grow xenograft tumors from immortalized cell lines, as well as, primary tumors from
patients, in an organism that can grow to a scale comparable to the size of humans.
Previous animal models have been limited due to small animal size, preventing the
development of tumors that are comparable in size and location to humans, and other
large animal models are less predictable and reproducible. With this controlled
immunocompromised model, the capability to engraft tumors would allow for more
relevant and efficient pre-clinical trials.
The growth of the tumors in the subcutaneous space in the pigs was notable
immediately after engraftment and maintained a relatively steady growth rate beyond
that point. This controlled growth in conjunction with histology, which showed no signs
of host immune rejection from the myeloid cells that remain in immunocompromised
conditions, indicate that this model is capable of supporting xenografts even with the
moderate cell injections of 1.2x106 used here. The current study stopped tumor growth
at 36 days as a pre-determined end point for consistency and direct comparisons with
the mouse models, not due to health effects or tumor burden, which commonly prevent
murine models from producing larger tumors. Beyond that, should this model be used
for subcutaneous treatments, the immunocompromised state of the pigs did not leave
them susceptible to increased systemic disease, as seen by their steady increase in
196
weight, lack of clinical symptoms, and their lack of metastases found on necropsy.
Future studies should be conducted in order to determine if orthotopic engraftment can
be supported by the animals without notable health effects.
Comparison of the porcine xenografted Panc01 tumors to the murine Pan02
tumors and PDAC murine xenografted PDX tumors, is based upon the latter two
model’s well established roll in studying cancer therapies.24,47,48 Based on histology, the
xenograft tumors also share comparable features of interest with accepted human-
tumor xenograft models. The PDX model is similar to the Pan02 model in that tumor
cells are subcutaneously injected into the mouse; however, the model differs in several
important ways (Fig 5A-B). The PDX tumors are grown in NSG, immunocompromised,
mice, and develop glandular structures that express CK19. These glands often have
distinct lumens and will often be filled with clear space, small amounts of secretory
product, inflammatory cells, and/or cellular debris. The most common type of pancreatic
tumor in humans is pancreatic ductal adenocarcinoma, which is histologically
characterized by polygonal cells often arranged into glandular structures. Similar to the
PDX models, the Panc01 tumors from the immunocompromised pigs also closely
resemble this microscopic morphology and are consistent with observations in human
patients. This includes cells with marked pleomorphism and prominent mitotic figures.
Another feature of the tumors in PDX mice is the fairly prominent fibrovascular stroma,
which is not present in the Pan02 tumor. However, the pig tumors do exhibit an
intermediate stroma again suggesting it is more similar to the PDX than the Pan02. The
tumor stroma seems to be a pretty important player and is often a barrier to treatment.49-
51 Additionally, agreeing with previous studies, we did not find ductal cell maker CK19
197
on our Pan02 tumors.52 On the other hand, the murine PDX and porcine Panc01 tumors
both express this marker, establishing their similarity to PDAC tumors.53 Thus, this
model recapitulates many features found in patient adenocarcinoma and is comparable
to PDX tumors, which are currently considered superior animal model for tumor
propagation and ex vivo studies.
For the feasibility studies described here, we used IRE as an example
technology that can benefit from tumors propagated in the RAG2/IL2RG pigs. Due to
electroporation-based therapies relying on exposing targeted cells to a specific electric
field, the conductivities of the tissues in the region of interest are critical to determining
treatment outcome. Thus, conductivity is one of the principal parameters affecting the
production of lesions due to irreversible electroporation, and preclinical animal models
must exhibit similar electrical signatures to their human counterparts to maximize
clinical translatability. Additionally, the morphology of cells affects the local electric
potential they experience when an external electric field is applied.54-56 Thus, faithful
recapitulations of human tumor tissue should be similar in terms of cell shape, size,
distribution, and intracellular content as this will ultimately affect the treatment outcome.
Notably, the PDX tumors presented in this study have been shown to recapitulate the
histological characteristics of human PDAC samples.24 Thus, the similar morphology of
cells in the Panc01 tumors introduced here indicates similarity to primary tumor samples
as well. Though the Panc01 tumors exhibited fewer ductal structures than the human
PDX tissue, this did not introduce differences in measured electrical properties. This
similarity is likely the result of the similar cell size and packing density between the PDX
and Panc01 tissue (Fig. 5). However, it is possible that the relatively large samples
198
(~0.2 cm3), excision process, and PBS wash introduce minute changes in tissue
structure/composition that limit the ability of our methods to resolve small spatial
differences in conductivity due to the presence of these ductal structures. Despite this,
we have shown that the bulk tissue response is similar between tissue models,
suggesting that the Panc01 platform exhibits a similar bulk response to applied fields as
currently established tissue surrogates.
To assess the electrical similarity of the subcutaneous Panc01 tumors to locally
advanced pancreatic cancer (LAPC) observed in humans, we conducted a series of
experiments in which electrical pulses were delivered across flat-plate electrodes to
determine electrical conductivity as a function of applied electric field. Our results
demonstrate that the Panc01 tumors grown in RAG2/IL2RG pigs have similar electrical
behavior to spontaneous tumors in humans, as well as murine-propagated human
PDAC tissue, increasing our confidence that the local electric field produced in the pig-
derived tumors will be similar to that which arises in human neoplasms.
Together, these data support the feasibility of the immunocompromised pig model
for evaluating the clinical relevance of novel medical devices and other surgical
procedures, including those that rely on accurate recapitulation of human tumor
morphology and electrical properties. Additionally, the anatomical features, including
size, of the RAG2/IL2RG pigs is more similar to that of humans compared to most
common pre-clinical animal models, making these swine ideal for prototype testing
without the need for a significant scale down. Here we have shown for the first time that
the dielectric properties of RAG2/IL2RG pigs are similar to those of humans. Overall,
these data indicate that this immunocompromised model is capable of recapitulating
199
physiologically relevant tumors comparable to other commonly utilized tumor models
with several novel advantages. Together, this suggests that this model will be highly
useful and capable of bridging the gap of translating ablation therapies from the bench
to clinical application.
5.7 Abbreviations
SCID: Severe combined immunodeficiency; IRE: irreversible electroporation; PDX:
patient derived xenograft
5.8 Funding Sources
This work was supported by the Virginia Maryland College of Veterinary Medicine
(I.C.A.); The Virginia Tech Institute for Critical Technology and Applied Sciences Center
for Engineered Health (I.C.A. and R.V.D.); The Virginia Biosciences Health Research
Corporation Catalyst (R.V.D.); The Focused Ultrasound Foundation (I.C.A.); The
National Institutes of Health R21OD027062 (K.L.), R21 R21EB028429 (I.C.A.),
R01CA213423 (R.V.D.), and P01CA207206 (R.V.D.). The content is solely the
responsibility of the authors and does not necessarily represent the official views of the
NIH or any other funding agency.
200
5.9 References
1 Howlader N, N. A., Krapcho M, Garshell J, Miller D, Altekruse SF, Kosary CL, Yu
M, Ruhl J, Tatalovich Z,Mariotto A, Lewis DR, Chen HS, Feuer EJ, Cronin KA
(eds). SEER Cancer Statistics Review, 1975-2012, National Cancer Institute.
Bethesda, MD. http://seer.cancer.gov/csr/1975_2012/, based on November 2014
SEER data submission, posted to the SEER web site, April 2015.
2 Riall, T. S. et al. Pancreatic cancer in the general population: improvements in
survival over the last decade. Journal of gastrointestinal surgery 10, 1212-1224
(2006).
3 Oettle, H. et al. Adjuvant chemotherapy with gemcitabine vs observation in
patients undergoing curative-intent resection of pancreatic cancer: a randomized
controlled trial. Jama 297, 267-277 (2007).
4 Conroy, T. et al. FOLFIRINOX versus gemcitabine for metastatic pancreatic
cancer. New England Journal of Medicine 364, 1817-1825 (2011).
5 Sultana, A. et al. Meta-analyses of chemotherapy for locally advanced and
metastatic pancreatic cancer. Journal of Clinical Oncology 25, 2607-2615 (2007).
6 Stocken, D. et al. Meta-analysis of randomised adjuvant therapy trials for
pancreatic cancer. British journal of cancer 92, 1372-1381 (2005).
7 Arcidiacono, P. G. et al. Feasibility and safety of EUS-guided cryothermal
ablation in patients with locally advanced pancreatic cancer. Gastrointestinal
endoscopy 76, 1142-1151 (2012).
201
8 Carrafiello, G. et al. Microwave ablation of pancreatic head cancer: safety and
efficacy. Journal of Vascular and Interventional Radiology 24, 1513-1520 (2013).
9 Girelli, R. et al. Feasibility and safety of radiofrequency ablation for locally
advanced pancreatic cancer. British Journal of Surgery 97, 220-225 (2010).
10 Vandamme, T. F. Use of rodents as models of human diseases. J Pharm
Bioallied Sci 6, 2-9, doi:10.4103/0975-7406.124301 (2014).
11 Tentler, J. J. et al. Patient-derived tumour xenografts as models for oncology
drug development. Nature reviews. Clinical oncology 9, 338-350,
doi:10.1038/nrclinonc.2012.61 (2012).
12 Consortium, M. G. S. Initial sequencing and comparative analysis of the mouse
genome. Nature 420, 520 (2002).
13 Rossant, J. & McKerlie, C. Mouse-based phenogenomics for modelling human
disease. Trends in molecular medicine 7, 502-507 (2001).
14 Paigen, K. A miracle enough: the power of mice. Nature medicine 1, 215 (1995).
15 De Jong, M. & Maina, T. Of mice and humans: are they the same?—Implications
in cancer translational research. Journal of Nuclear Medicine 51, 501-504 (2010).
16 Perlman, R. L. Mouse models of human disease: An evolutionary perspective.
Evol Med Public Health 2016, 170-176, doi:10.1093/emph/eow014 (2016).
17 Sabounchi, N. S., Rahmandad, H. & Ammerman, A. Best-fitting prediction
equations for basal metabolic rate: informing obesity interventions in diverse
populations. Int J Obes (Lond) 37, 1364-1370, doi:10.1038/ijo.2012.218 (2013).
18 Blagosklonny, M. V. Big mice die young but large animals live longer. Aging
(Albany NY) 5, 227-233, doi:10.18632/aging.100551 (2013).
202
19 Balmain, A. & C. Harris, C. Carcinogenesis in mouse and human cells: parallels
and paradoxes. Carcinogenesis 21, 371-377 (2000).
20 Greenberg, R. A., Allsopp, R. C., Chin, L., Morin, G. B. & DePinho, R. A.
Expression of mouse telomerase reverse transcriptase during development,
differentiation and proliferation. Oncogene 16, 1723 (1998).
21 Smith, V., Wirth, G. J., Fiebig, H. H. & Burger, A. M. Tissue microarrays of
human tumor xenografts: characterization of proteins involved in migration and
angiogenesis for applications in the development of targeted anticancer agents.
Cancer genomics & proteomics 5, 263-273 (2008).
22 Garrido-Laguna, I. et al. Tumor engraftment in nude mice and enrichment in
stroma- related gene pathways predict poor survival and resistance to
gemcitabine in patients with pancreatic cancer. Clinical cancer research : an
official journal of the American Association for Cancer Research 17, 5793-5800,
doi:10.1158/1078-0432.Ccr-11-0341 (2011).
23 Tratar, U. L., Horvat, S. & Cemazar, M. Transgenic mouse models in cancer
research. Frontiers in oncology 8 (2018).
24 Brock, R. M. et al. Patient derived xenografts expand human primary pancreatic
tumor tissue availability for ex vivo irreversible electroporation testing. Frontiers
in Oncology 10 (2020).
25 Sieren, J. C. et al. Development and translational imaging of a TP53 porcine
tumorigenesis model. The Journal of clinical investigation 124, 4052-4066
(2014).
203
26 Ganderup, N. C., Harvey, W., Mortensen, J. T. & Harrouk, W. The minipig as
nonrodent species in toxicology—where are we now? International journal of
toxicology 31, 507-528 (2012).
27 Swindle, M. & Smith, A. Swine in the Laboratory: Surgery. Anesthesia, Imaging,
and Experimental Techniques, 40-42 (2007).
28 Swindle, M., Makin, A., Herron, A., Clubb Jr, F. & Frazier, K. Swine as models in
biomedical research and toxicology testing. Veterinary pathology 49, 344-356
(2012).
29 Segatto, N. V. et al. The oncopig cancer model as a complementary tool for
phenotypic drug discovery. Frontiers in pharmacology 8, 894 (2017).
30 Maher, B. Tissue engineering: How to build a heart. Nature News 499, 20 (2013).
31 Crile, G. W., Hosmer, H. R. & Rowland, A. F. The electrical conductivity of animal
tissues under normal and pathological conditions. American Journal of
Physiology-Legacy Content 60, 59-106 (1922).
32 Davalos, R. V., Mir, L. M. & Rubinsky, B. Tissue ablation with irreversible
electroporation. Ann Biomed Eng 33, 223-231 (2005).
33 Maor, E., Ivorra, A., Leor, J. & Rubinsky, B. The effect of irreversible
electroporation on blood vessels. Technol Cancer Res Treat 6, 307-312 (2007).
34 Garcia, P. A. et al. Non-thermal irreversible electroporation (N-TIRE) and
adjuvant fractionated radiotherapeutic multimodal therapy for intracranial
malignant glioma in a canine patient. Technol Cancer Res Treat 10, 73-83
(2011).
204
35 Neal, R. E., 2nd et al. Successful treatment of a large soft tissue sarcoma with
irreversible electroporation. Journal of clinical oncology : official journal of the
American Society of Clinical Oncology 29, e372-377,
doi:10.1200/JCO.2010.33.0902 (2011).
36 Li, W., Fan, Q., Ji, Z., Qiu, X. & Li, Z. The Effects of Irreversible Electroporation
(IRE) on Nerves. PLOS ONE 6, e18831, doi:10.1371/journal.pone.0018831
(2011).
37 Lee, K. et al. Engraftment of human iPS cells and allogeneic porcine cells into
pigs with inactivated RAG2 and accompanying severe combined
immunodeficiency. Proc Natl Acad Sci U S A 111, 7260-7265,
doi:10.1073/pnas.1406376111 (2014).
38 Choi, Y. J. et al. Recombination activating gene-2(null) severe combined
immunodeficient pigs and mice engraft human induced pluripotent stem cells
differently. Oncotarget 8, 69398-69407, doi:10.18632/oncotarget.20626 (2017).
39 Choi, Y. J. et al. Partial loss of interleukin 2 receptor gamma function in pigs
provides mechanistic insights for the study of human immunodeficiency
syndrome. Oncotarget 7, 50914-50926, doi:10.18632/oncotarget.10812 (2016).
40 Kang, J. T. et al. Biallelic modification of IL2RG leads to severe combined
immunodeficiency in pigs. Reprod Biol Endocrinol 14, 74, doi:10.1186/s12958-
016-0206-5 (2016).
41 Lei, S. et al. Increased and prolonged human norovirus infection in RAG2/IL2RG
deficient gnotobiotic pigs with severe combined immunodeficiency. Sci Rep 6,
25222, doi:10.1038/srep25222 (2016).
205
42 Lei, S. et al. Increased and prolonged human norovirus infection in RAG2/IL2RG
deficient gnotobiotic pigs with severe combined immunodeficiency. Scientific
reports 6, 25222 (2016).
43 Lee, K. et al. Engraftment of human iPS cells and allogeneic porcine cells into
pigs with inactivated RAG2 and accompanying severe combined
immunodeficiency. Proceedings of the National Academy of Sciences 111, 7260-
7265 (2014).
44 Basel, M. T. et al. Human xenografts are not rejected in a naturally occurring
immunodeficient porcine line: a human tumor model in pigs. BioResearch open
access 1, 63-68 (2012).
45 Jain, R., Fischer, S., Serra, S. & Chetty, R. The use of cytokeratin 19 (CK19)
immunohistochemistry in lesions of the pancreas, gastrointestinal tract, and liver.
Applied Immunohistochemistry & Molecular Morphology 18, 9-15 (2010).
46 Halter, R. J., Schned, A., Heaney, J., Hartov, A. & Paulsen, K. D. Electrical
properties of prostatic tissues: I. Single frequency admittivity properties. the
Journal of Urology 182, 1600-1607 (2009).
47 Guo, S. et al. Enhanced Electric Pulse Technology for the Ablation of Pancreatic
Cancer. Advances in Pancreatic Cancer, 115 (2018).
48 Moschidis, A. et al. Gemcitabine in combination with Oxaliplatin in Pancreatic
(PAN-02) tumor bearing mice. Journal of Clinical Oncology 22, 4153-4153
(2004).
49 Hofmeister, V., Schrama, D. & Becker, J. C. Anti-cancer therapies targeting the
tumor stroma. Cancer Immunology, Immunotherapy 57, 1-17 (2008).
206
50 Ma, W. et al. Tumor-stroma ratio is an independent predictor for survival in
esophageal squamous cell carcinoma. Journal of Thoracic Oncology 7, 1457-
1461 (2012).
51 Dittmer, J. & Leyh, B. in Seminars in cancer biology. 3-15 (Elsevier).
52 Torres, M. P. et al. Novel pancreatic cancer cell lines derived from genetically
engineered mouse models of spontaneous pancreatic adenocarcinoma:
applications in diagnosis and therapy. PloS one 8, e80580 (2013).
53 Zhou, R., Xu, X., Liu, M., Wu, X. & Li, R. Immunophenotypes of ductal epithelial
cells in advanced pancreatic ductal adenocarcinoma. Digestion 99, 247-251
(2019).
54 Ivey, J. W. et al. Targeted cellular ablation based on the morphology of malignant
cells. Scientific reports 5, 17157 (2015).
55 Maor, E., Ivorra, A. & Rubinsky, B. Non thermal irreversible electroporation: novel
technology for vascular smooth muscle cells ablation. PloS one 4, e4757 (2009).
56 Murovec, T., Sweeney, D. C., Latouche, E., Davalos, R. V. & Brosseau, C.
Modeling of transmembrane potential in realistic multicellular structures before
electroporation. Biophysical journal 111, 2286-2295 (2016).
57 Lee, K. et al. Piglets produced from cloned blastocysts cultured in vitro with
GM‐CSF. Molecular reproduction and development 80, 145-154 (2013).
58 Yuan, L., Jobst, P. M. & Weiss, M. in Gnotobiotics 349-368 (Elsevier, 2017).
59 Abcam. (Abcam Corporation Cambridge, 2005).
207
60 Bhonsle, S. et al. Characterization of irreversible electroporation ablation with a
validated perfused organ model. Journal of Vascular and Interventional
Radiology 27, 1913-1922. e1912 (2016).
61 Bhonsle, S., Lorenzo, M. F., Safaai-Jazi, A. & Davalos, R. V. Characterization of
Nonlinearity and Dispersion in Tissue Impedance During High-Frequency
Electroporation. IEEE Transactions on Biomedical Engineering 65, 2190-2201
(2017).
62 Beitel-White, N., Bhonsle, S., Martin, R. & Davalos, R. V. in 2018 40th Annual
International Conference of the IEEE Engineering in Medicine and Biology
Society (EMBC). 4170-4173 (IEEE).
63 Geddes, L. A. & Baker, L. E. The specific resistance of biological material—a
compendium of data for the biomedical engineer and physiologist. Medical and
biological engineering 5, 271-293 (1967).
64 Clark, D., Greenwell, J., Harper, A., Sankey, A. M. & Scratcherd, T. The electrical
properties of resting and secreting pancreas. The Journal of physiology 189, 247-
260 (1967).
65 Duck, F. A. Physical properties of tissues: a comprehensive reference book.
(Academic press, 2013).
66 McCann, H., Pisano, G. & Beltrachini, L. Variation in reported human head tissue
electrical conductivity values. Brain topography 32, 825-858 (2019).
67 Stoy, R. D., Foster, K. R. & Schwan, H. P. Dielectric properties of mammalian
tissues from 0.1 to 100 MHz; a summary of recent data. Physics in Medicine &
Biology 27, 501 (1982).
208
68 Neal, R. E. et al. In vivo characterization and numerical simulation of prostate
properties for non‐thermal irreversible electroporation ablation. The Prostate 74,
458-468 (2014).
209
5.10 Supplemental Figure
Figure 5-9 Supplemental Figure 5.S1. Genotyping of piglets carrying modified IL2RG and RAG2 gene. Six piglets were born and analyzed. During PCR analysis, RAG2 was not amplified from the Piglet #2 DNA sample, suggesting a large deletion. No wild-type allele was identified.
210
Chapter 6. Feasibility of Histotripsy Pancreas Ablation in a Large Pig Model
Alissa Hendricks-Wenger, Jessica Gannon, Neha Singh, Lauren Arnold, Michael Edwards, Khan Mohammad Imran, Margaret Nagai-Singer, Benjamin Tintera, Hannah Sheppard, Joan Vidal-Jove, David Luyimbazi, Mishal Mendiratto-Lala, Martha Larson,
Sheryl Coutermarsh-Ott, Irving C Allen, Eli Vlaisavljevich
This chapter is excerpted from a manuscript in preparation for submission to Ultrasound in Medicine and Biology.
211
6.1 Abstracts
Pancreatic cancer is one of the most lethal forms of cancer, with a 5 year-survival rate
of 9%. To address this, many new therapeutic modalities are being developed. One
modality is the focused-ultrasound therapy Histotripsy, a non-invasive, non-ionizing, and
non-thermal ablation modality that has been shown to be effective in treating hepatic
malignancies. However, when translating an ultrasonic modality to the pancreas there
are limitations that need to be considered, including 1) achieving an acoustic window
and 2) generating an ablation lesion without causing moderate or severe complications.
For this study, we used healthy pigs in order to test the efficacy of our custom 32
element, 500kHz histotripsy transducer that is comparable to human scale devices. To
obtain an ideal acoustic window, a pilot study was conducted using pigs that had been
fasted for at least 12 hours. While we were able to use our system to generate a bubble
cloud and lesion within the liver, we were unable to get an acoustic window for targeting
the pancreas. Using a laxative and simethicone custard in combination with fasting, the
gas within the porcine gastrointestinal system was reduced. This allowed for easier
ultrasound targeting of the pancreas, generation of a bubble cloud within the pancreas,
and creation of a clear lesion. Acute necropsy demonstrated mild bruising to intestines
and the stomach that was near the focus as well as in some loops of pre-focal bowel.
Following a group (n=4) for 1-week post treatment through behavioral observations,
diet, and blood work, we did not find signs of any complications. At 1 week on contrast
enhanced CT imaging and necropsy, there were no signs of bruising or abnormalities
212
on any organs. Overall, these results suggest that our histotripsy system and degassing
protocol creates the opportunity to target the porcine pancreas in future studies.
6.2 Keywords:
Pancreas, Pancreatic Cancer, Histotripsy, Porcine, Ultrasound Imaging
213
6.3 Introduction
Pancreatic cancer is considered a fatal malignancy with a 5-year survival rate lower
than 9% and the majority of patients dying within 6 months of diagnosis [1]. Most
complications that contribute to this disease are associated with a late diagnosis, rapid
local progression with vascular compromise, and metastasis. The current gold standard
for treatment includes surgery [2]. However, the location of the tumor is a strong
prognostic factor, and its proximity to vital vascular structures and local infiltration into
the adjacent mesenteric fat are major limitations precluding resection in a large number
of patients [3]. Despite improved outcomes with current frontline therapeutics, a high
incidence of refractory disease from chemo- and radio-resistance, as well as local and
distant recurrence, still pose significant challenges [2]. There are a multitude of tumor
ablation methods including radiofrequency (RFA), microwave (MWA), cryoablation, high
intensity focused ultrasound (HIFU), and irreversible electroporation (IRE) [4-6].
However, only HIFU and IRE have been developed as stand-alone or combination
therapeutic options for pancreatic cancer, given the high complication risk and technical
challenges of RFA and MWA [7, 8].
IRE and HIFU have shown some improvement over standard of care in terms of
morbidity, tissue conservation, improved surgical accessibility, and decreased
hospitalization [9, 10]. However, these strategies can be associated with significant
complications, and many patients are not candidates due to local tumor progression,
tumor size, and location near critical structures [10, 11]. HIFU, among other thermal
214
ablation methods, runs a high risk for thermal injury to critical structures near the
ablation zone, particularly pancreatic duct injury. This damage can result in a leak and
severe pancreatitis or even stenosis with resultant obstructive pancreatitis [9].
Additionally, the lack of real-time treatment feedback limits the ability to create precise
ablation zones. The invasive needle puncture used in IRE, inability to treat large (>3 cm
diameter) or multiple tumor nodules, and incomplete ablation near major vessels are
additional limitations. All of these limitations render these treatment modalities as
challenging for clinical use [9, 10]. As such, there remains an unmet need for a new
ablation method that can address the limitations of currently available ablation methods
for pancreatic cancer.
Histotripsy is a non-invasive, non-ionizing, and non-thermal focused ultrasound
ablation method that destroys tissue through the precise control of acoustic cavitation
[12, 13]. Unlike thermal HIFU, histotripsy disrupts tissue through mechanical effects
(i.e., cavitation) [13]. Using high-pressure pulses applied by an external transducer, a
cavitation “bubble cloud” is generated at the target [13, 14], and the expansion and
collapse of the bubbles ablates the tissue into an acellular homogenate [13, 14]. Since
histotripsy is non-thermal, it does not have the many limitations associated with thermal
ablation [15, 16]. Histotripsy can produce consistent and rapid ablation, even in close
proximity to major vessels, with millimeter precision and well-defined boundaries
(<2mm) [17]. The high accuracy of histotripsy is a direct result of the non-thermal
mechanism, which results in a “binary” treatment outcome with sharp boundaries and
millimeter precision. This is a significant improvement upon thermal ablation methods
that do not have well-defined margins and are often surrounded by a centimeter or more
215
of partially treated tissue due to thermal spread [17, 18]. Furthermore, histotripsy can
preserve critical structures such as large vessels and bile ducts due to the higher
mechanical strength of these structures [17, 18]. Additional benefits include real-time
imaging feedback [19] and the ability to cover tumor nodules of arbitrary size and shape
[20]. Due to these features, histotripsy has the potential to overcome the weaknesses of
current treatment methods and benefit many diverse diseases [21].
Previous in vivo animal studies have investigated histotripsy’s use for precise and
complete liver ablation. These studies also showed that histotripsy can generate tissue
selective ablation in which the target tissue is completely destroyed into an acellular
homogenate, while critical structures (vessels and bile ducts) are preserved [17]. These
initial studies led to the development of a clinical prototype histotripsy system, which
was subsequently tested in a pig model, to show the ability of producing clinically
relevant ablation zones in livers that are closer to human size. Additionally, these
preclinical results led to a Phase I clinical trial in the FUS Ablation Oncology Unit of
Hospital University Mutua Terrassa (Barcelona, Spain), led by Dr. Joan Vidal Jove, one
of our key clinical collaborators. Favorable results from the trial demonstrated that
histotripsy can non-invasively produce well-tolerated treatments and rapid involution of
ablation zones without significant device-related adverse events [22].
Previous work has shown the feasibility of focused ultrasound (FUS) for targeting the
pancreas. For example, a series of clinical studies have been performed on non-
resectable pancreatic tumors (Stages III/IV) in the FUS Ablation Oncology Unit of
Hospital University Mutua Terrassa in Barcelona, Spain [22]. Results showed a
sufficient acoustic window available to apply HIFU to pancreatic tumors under
216
ultrasound guidance. Clinical responses (thermal ablation achieved) were 82% in all
cases, sustained at 8 weeks of the procedure, with 12 complete responses (21% of
patients, 10 stage III, 2 stage IV) at the end of a combined HIFU and chemotherapy
regime. However, results also showed major complications due to the thermal
mechanism that included pancreatitis with GI bleeding and skin burning grade III that
required plastic surgery. Overall, these results showed that FUS can target the
pancreas non-invasively under ultrasound guidance with the potential to achieve
effective treatments outcomes. Results also showed limitations of HIFU due to the
thermal mechanism, including injury to critical structures and overlying tissue. We
expect histotripsy to overcome these limitations due to its non-thermal mechanism,
allowing for safer, more precise, and more effective pancreatic ablation.
While prior studies highlight the potential of histotripsy for non-invasive tumor
ablation in other organs, significant work remains to establish histotripsy’s potential in
pancreatic tumor ablation. The pancreas poses unique challenges for ablation
modalities due to the tissue characteristics and risk of damage to nearby critical
structures. In this study, we investigate the feasibility of histotripsy in the treatment of
healthy pancreatic tissue in an in vivo pig model.
6.4 Materials and Methods
6.4.1 Animal Monitoring and Anesthesia
217
Male pigs (6-8 weeks old, 5-8kg; Virginia Tech Swine Farm, Blacksburg, VA)
were monitored three times during their first week to acclimate to their new housing, and
then daily after treatment in accordance with Virginia Tech IACUC protocols. At each
check, general health was monitored looking for changes in behavior, diet, and skin
lesions. Blood was collected into green and lavender vacutainer tubes at baseline and
post-treatment days 1 and 5. Tubes were submitted to the Virginia Maryland Animal
Laboratory Services (Blacksburg, VA) for analysis for complete blood cell and
pancreatic enzyme panels. General workflow shown in Figure 1.
For treatment, animals were sedated by using an intramuscular injection of the
cocktail of Telazol, Ketamine, and Xylazine administered in intervals approximately 15-
20 minutes apart, or when the animal began to demonstrate increased muscle tone.
Animals’ heart rate, blood oxygen levels, respiratory rate, and temperature were
monitored during treatment by trained personnel. Until fully recovered, animals were
visually monitored for 2-3 hours in recovery kennels before being returned to their
standard housing.
Figure 6-1Study timeline demonstrating the relative time for treatment, custard feeding, blood draws, and CT imaging.
218
6.4.2 Histotripsy Systems and Pressure Calibrations
Histotripsy was performed in vivo using a custom 32-element, 500 kHz large
animal histotripsy transducer with a geometric focus of 78 mm, an elevational aperture
size of 112 mm, and a transverse aperture size of 128 mm. Corresponding f-numbers
include 0.61 (transverse) and 0.70 (elevational). To generate short therapy pulses <2
cycles, the transducer was controlled via a custom high-voltage pulser with a field-
Figure 6-2 HistoSonics cart with 500 kHz transducer and coaxial US imaging probe mounted onto a micropositioner (A). Freehand imaging by radiologist to identify acoustic window (B). UMC bowl with therapy transducer submerged via adjustable arm (C).
219
programmable gate array (FPGA) board (Altera DE0-Nano Terasic Technology, Dover,
DE, USA) programmed for histotripsy therapy pulsing. The transducer was mounted
onto a prototype clinical histotripsy treatment cart (HistoSonics, Ann Arbor, MI, USA)
consisting of an ultrasound imaging system, robotic micro-positioner, and customized
software to create volumetric ablations with histotripsy. A central hole through the
therapy transducer allowed a 3 MHz curvilinear imaging probe (Model C5-2, Analogic
Corp., Peabody, MA) to be coaxially aligned to enable real-time treatment guidance and
monitoring. [24, 25]. During histotripsy treatments, the transducer was triggered from
the HistoSonics software while being powered by a high voltage DC power supply
(GENH750W, TDK-Lambda), and the system was controlled using a custom user-
interface operated through MATLAB (MathWorks). The overall setup for the treatment is
illustrated in Figure 2A.
6.4.3 Hydrophone Focal Pressure Measurements and Beam Profiles
Focal pressure waveforms for the 500 kHz transducer were collected using a high
sensitivity rod hydrophone (HNR-0500, Onda Corporation, Sunnyvale, CA, USA) and a
custom-built fiber optic probe hydrophone (FOPH) [23, 24] in degassed water at the
focus of the transducer. 1D focal beam profiles of the transducer were measured in the
lateral, elevational, and axial directions with the rod hydrophone at a peak negative
pressure (p-) of ~1.8 MPa. The FOPH directly measured focal pressures up to a p- limit
of ~20 MPa due to cavitation on the tip of the FOPH fiber. Pressure waveforms at higher
p- were then estimated by a linear extrapolation [25]. All waveforms were captured at a
220
sample rate of 500MS/s using a Tektronix TBS2000 series oscilloscope with the
waveforms averaged over 128 pulses and recorded in MATLAB. This procedure with is
described in previous studies further details [25, 26].
6.4.4 Histotripsy In Vivo Pancreas Ablation Procedure
The feasibility of histotripsy for pancreas ablation was tested in vivo using
healthy, young pig. This custom system, which has been used in previous large animal
in vivo histotripsy studies [27-29], consists of the 500 kHz histotripsy transducer with co-
axially aligned ultrasound imaging connected to an articulating arm with 4 degrees of
freedom (Company Source of Arm). Six male pigs were purchased at 6 weeks of age
from the pig farm at Virginia Tech. Once received, animals were monitored three times
during their first week to acclimate to their new holding pins, and then daily after
treatment, in accordance with Virginia Tech IACUC protocols. At each check, general
health was monitored, and blood was collected into vacutainer tubes at baseline and
post-treatment day 5.
For the 4 days leading to treatments, the animals were fed a sweetened custard
once daily with simethicone (2 mL/pig) and bisacodyl (0.3 mg/kg) in order to minimize
the intestinal contents and gas present on the day of treatment. Pigs were fasted from
food after their evening meal the day before treatment, and were given an additional
custard 2 hours before the first treatment of the morning. Immediately before treatment,
hair was removed over the abdomen by shaving with electric razor and then Nair
(Naircare, Ewing, NJ, USA) was applied for 10 minutes before being removed by wiping
221
off with a wet towel. For treatment, pigs were anesthetized with an intramuscular
injection of the cocktail TKX (Telazol-Ketamine-Xylazine) and placed in the dorsal
recumbent position on the treatment table with feet loosely restrained to maintain a
stable position (Fig.2B). Heart rate, blood oxygen levels, and temperature were
monitored for all pigs during treatment by trained personnel.
Before treatment, freehand ultrasound imaging was performed by a board-
certified veterinary radiology (M.L.) or veterinary radiology resident (M.E.) to identify the
pancreas and determine the desired treatment location using both a linear ultrasound
imaging probe with a frequency range of 10-18 MHz (L18-10L30H-4, Telemed,
Lithuania, EU) and a 3 MHz curvilinear imaging probe (Model C5-2, Analogic Corp.,
Peabody, MA) (Fig.2B). Custom Ioban (3M) drapes with a hole large enough to
establish an acoustic window were attached to the pig’s abdomen. A plastic ultrasound
mediated coupling (UMC) bowl, with the base removed attached to an articulating arm,
was mounted to the surgical table. The bowl was then manually adjusted with the
articulating arm to align the hole of the bowl with the acoustic treatment window. Excess
drape was carefully pulled into the bowl and clamped to the upper rim. Before
treatment, a water tub was filled with water and degassed for ~2 hours, with this water
being used to fill the UMC bowl for ultrasound coupling. The therapy transducer and
coaxial imaging probe were mounted onto a mechanical arm containing a
micropositioner (Histosonics Inc., Ann Arbor, MI) [28], was manually positioned to
aligned with the acoustic window identified during freehand imaging (Fig.2C). A bubble
cloud was generated in degassed water prior to treatment to align the focus of the
transducer with system’s imaging screen
222
For treatment, the coaxial imaging probe was used to align the focus of the
transducer to the targeted location. The transducer was then turned on and histotripsy
was applied to the target using single cycle pulses and a pulse repetition frequency
(PRF) of 500Hz. The cavitation threshold within the tissue was determined by slowly
increasing the pressure until a robust bubble cloud was generated. The transducer
swept through a 1 – 1.5 cm diameter spherical ablation volume delivering 500 pulses
per treatment point with a peak negative pressure of ~21 MPa. Throughout the entire
procedure, ultrasound imaging was used to monitor the treatment in real-time. To
ensure a complete ablation of the targeted pancreas, a second treatment to the exact
same volume was immediately done after the first treatment. After treatment, the
therapy transducer was removed and freehand ultrasound imaging was performed
assess for any tissue damage. Animals in the acute group (n=2) were immediately
euthanized by an intracardiac injection of pentobarbital sodium and phenytoin sodium
(Euthasol) according to protocol. Animals in the 1-week group were anesthetized and
euthanized after CT. A full necropsy was conducted post-treatment by a board-certified
veterinary pathologist (S.C.O.).
6.4.5 Computed Tomography Imaging
Anesthetized pigs were placed in a supine position in a veterinary CT scanner
(Toshiba Aquilion 16-slice) for 1 week post treatment imaging. CTs were performed 1
week after treatment, based on prior studies that show that moderate and severe post-
223
operative complications occur in this window [30]. Images were analyzed by a board-
certified veterinary radiologist (M.L.) and veterinary radiology resident (M.E.).
6.4.6 Necropsy & Histological Analysis
The acute and 1-week-survival groups animals’ necropsy and tissue harvest was
preformed either immediately and 1-week post-treatment, respectively, by a board-
certified veterinary pathologist (S.C.O.). Gross damage to internal organs was
determined with inspection in situ and ex vivo with bread slices spaced approximately
1cm apart. Bread slicing of pancreatic tissue was performed after fixation due to the
friable nature of the pancreatic tissue. Pancreatic tissue and any tissue with noted gross
changes were visually inspected and fixed in 10% formalin for at least 24 hours before
sectioning, embedding, and staining. All tissues were stained using hematoxylin and
eosin (H&E) to assess for histotripsy damage to the tissue parenchyma and any other
changes in structure and density of collagen and other tissue structures within the
ablation volume. Analysis was led by a board-certified veterinary pathologist (S.C.O.) to
ensure accuracy and avoid biases.
6.5 Results
6.5.1 Establishing Safe, Non-Invasive Therapeutic Protocol
Targeting the liver with our histotripsy system clearly showed a describable bubble
cloud (Fig.3A) and on gross necropsy showed a clear ablation region (Fig.3B).
224
However, there was a more difficult time getting clear ultrasound images of the bubble
cloud within the pancreas. In the pilot group, in which the pigs were only fasted for ~12
hours before treatment, there was difficulty imaging the pancreas with the off-set
imaging (Fig.3C). When the pigs were fed the laxative, simethicone custard on the
prescribed regime, the dynamic nature of the bubble cloud was visible on ultrasound.
Due to the hyperechoic nature of the pancreas relative to the bubble cloud, it is difficult
to discern the bubble cloud in still frame images of the pancreas (Fig.3D).
Figure 6-3 Bubble cloud formed in liver on US imaging from HistoSonics system (pig 1, 8-6)(A). No bubble cloud visualized on US imaging and did not clearly identify the pancreas without pudding (pig 6, 8-13)(B). Bubble cloud generated inside pancreas where pig was fed pudding prior to histotripsy treatment (28-53) (C).
225
Intestinal gas interference during treatment prevented ideal imaging quality and
bubble formation (Fig.3C) leading to pre-focal intestinal bruising (Fig.4A-B). Bruising
was more highly concentrated in the more gaseous large intestine, indicated by the
green arrow, and near the targeted region, indicated by the blue arrows. Post treatment
H&E histopathology indicated that from a sample of the bruised colon, the muscularis
Figure 6-4 Intestinal gas interference with treatment prevented ideal imaging quality and bubble formation during treatment leading to pre-focal intestinal bruising (A-B) more highly concentrated in the more gaseous large intestine, indicated by the green arrow, and near the targeted region, indicated by the blue arrows. Post treatment H&E histopathology indicated that from a sample of the bruised colon, the muscularis was not damaged but there was a large amount of ablation within the submucosa (C-D).
226
was not damaged but there was a large amount of ablation within the submucosa
(Fig.4C-D).
When using the custard as part of the bowel preparation, the improved imaging
(Fig.5A) allowed for improved ablation. There was evident lesion generation within the
pancreas within the laxative, simethicone custard group of pigs. Post-treatment US
imaging (Fig.5B) clearly shows a hypoechoic region, consistent with a histotripsy
ablation, in the same target location. In these animals, while we did have off target
effects such as bruising to the intestinal tissues (Fig.6), this was reduced from the pilot
group that was not fed the custard mix.
6.5.2 Physiological Outcomes
All animals included in the 1 week-survival group (n=4) recovered well from
treatment, and were returned to standard housing with their normal, unrestricted diet
within 4 hours of treatment. Over the week following treatment, the animals had no
signs of debilitating side effects. Specifically, there was no note of: fatigue, pain,
vomiting, diarrhea, or discolored stool. All treated animals maintained normal weight,
Figure 6-5 US imaging of pancreas before histotripsy (A) and after treatment with clear hypoechoic region identifying the targeted ablated volume (B). Dotted red circle indicated the approximate region treated
227
eating and drinking habits, levels of activity, and socialization. WBC, amylase, and
lipase levels of all animals were unaffected by histotripsy treatment 1 and 5 days post-
treatment compared to their baseline values determined 5 days prior to treatment
(Fig.7). No abnormalities were noted on the complete blood cell panel. From the 1-week
post treatment group, CT showed that there was no free fluid in the abdomen (Fig.8).
Chronic necropsy demonstrated that there was no gross damage to any organ in situ
and there was no free fluid or intestinal contents in the abdomen (Fig.9). There was no
splenomegaly, and bread slices to liver and spleen tissues further confirmed no off-
target ablation within those organs. Bread slices of formalin fixed pancreatic tissues did
not reveal any gross lesions. Histology from the 1-week necropsy showed no notable
lesions, hemorrhage, or inflammation within the pancreas (Fig.10).
Figure 6-6 With laxative, simethicone, and custard pretreatment protocol, there was signifcatly less bruising in preforcal bowel (A) indicated by the green arrow. Bruising around the treatment area was more concentrated than on the bowel, but was still less intense than the acute group (B), bruising indicated by blue arrow.
228
0 5 10
0
10
20
30
40
50
White Blood Cell
Days Post Arrival
WB
Cs
(x
10^
3 c
ell
s/u
L) Treatment
0 5 10
0
500
1000
1500
2000
Amylase
Days Post Arrival
Am
yla
se (
U/L
)
Treatment
0 5 10
0
5
10
15
20
Lipase
Days Post Arrival
Lip
as
e (
U/L
)
Treatment Pig 1
Pig 2
Pig 3
Pig 4
Figure 6-8 White blood cell, amylase, and lipase serum levels for 1-week survival animals.
Figure 6-7 CT taken in the dorsal recumbent position (A) one week post treatment shows no free fluid in the peritoneal cavity (B)
229
Figure 6-10 Necropsy reviled no gross damage within the abdomen in situ (A-B). When excised in tota no damage was found on the exterior of the liver (C-D), the spleen (E), nor the pancreas (F-G).
Figure 6-9 1-week post treatment there was no signs of morbidity within the pancreatic tissue. Images representative of two animals.
230
6.6 Discussions
In this study, we investigated the feasibility of using histotripsy for pancreatic tissue
ablation in an in vivo porcine model. Targeting intraabdominal organs, such as the
pancreas, with ultrasonic imaging and therapies requires additional planning and
investigation to determine the ideal preoperative and operative procedures that can
minimize the effects of pre-focal, acoustically opaque gas pockets within the
gastrointestinal tract. Given that the two notable benefits of histotripsy as an ablation
modality are that it is non-invasive as well as ultrasound imaging guided, the focus of
this study was to determine the feasibility of this approach for targeting the pancreas. A
prior porcine study utilizing HIFU to treat the pancreas was able to perform the
procedure without the use of nasogastric tube or other more invasive technique to
remove the gas from the gastrointestinal tract [31]. However, given HIFU’s thermal
mechanism of ablation, they were limited to using MR imaging for planning due to the
lack of ultrasonic methods for measuring ablative temperature change. In this study, a
combination of simethicone, laxative, and custard, previously reported non-invasive
methods for degassing the stomach and intestines [31], allowed for histotripsy to be
applied solely with ultrasound guidance (Fig.5).
Potential off-target effects are a significant hurdle for therapeutic ablation techniques
like histotripsy, particularly for structures like the pancreas which are surrounded by
critical structures. Results from this study suggest that histotripsy can be used to
effectively target the pancreas without causing damage to nearby, critical structures.
While off-target intestinal bruising was identified (Fig.6), there were no signs of
231
pancreatitis (Fig.7), intestinal perforation, intraabdominal bleeding, or pancreatic fistulas
(Fig.8 and Fig.9). Determining if an ablation therapy causes pancreatitis is important
due to the need to treat margins without causing negative side effects. Given that the
porcine pancreas is similar in anatomy and presentation of acute pancreatitis to humans
[32-34], they make an appropriate model for this style of investigation. A previous
porcine study testing the use of IRE within the pancreas showed a rise in WBCs,
amylase, and lipase as well as anorexia after treatment [35]. While this pilot study for
using histotripsy to target the pancreas is preliminary for targeting the pancreas, the
safety profile is evident. By being non-invasive, we did not risk infection. By propagating
the therapeutic effect directly to the targeted region, we did not agitate additional
pancreatic stroma to the point of causing acute pancreatitis.
In addition to the concern of pancreatitis previously reported in association with other
ablation modalities, there is an additional concern that histotripsy could destabilize the
mechanical structure of the pancreas resulting in a pancreatic fistula. The 2016
International Study Group of Pancreatic Fistula redefined a pancreatic fistula as post-
operative drainage from the pancreas that results in a sequelae that requires clinical
intervention to prevent mortality [36, 37]. From the 4 pigs that were treated with
histotripsy in the study and survived for 1 week to determine post-treatment side effects,
there were no changes to behavior or diet and no signs on CT (Fig.8) or necropsy
(Fig.9) to indicate that a pancreatic fistula was formed.
232
6.7 Conclusion
This study demonstrates the feasibility of histotripsy for non-invasive ablation within
the pancreas. Overall, this study demonstrated that histotripsy can be used to treat
targets within the pancreas without causing moderate nor severe side effects. Future
work will aim the further optimize histotripsy for pancreas ablation and determine the
potential of histotripsy to ablate pancreatic tumors.
6.8 Acknowledgements
Thanks for Kristen Eden, DVM, PhD for providing additional necropsy support for 2
animals when S.C.O. was unavailable.
233
6.9 References
[1] R. L. Siegel, K. D. Miller, and A. Jemal, "Cancer statistics, 2020," CA: a cancer
journal for clinicians, vol. 70, no. 1, pp. 7-30, 2020.
[2] E. G. Chiorean and A. L. Coveler, "Pancreatic cancer: optimizing treatment
options, new, and emerging targeted therapies," Drug design, development and
therapy, vol. 9, pp. 3529-45, 2015, doi: 10.2147/DDDT.S60328.
[3] A. Artinyan, P. A. Soriano, C. Prendergast, T. Low, J. D. Ellenhorn, and J. Kim,
"The anatomic location of pancreatic cancer is a prognostic factor for survival,"
HPB : the official journal of the International Hepato Pancreato Biliary
Association, vol. 10, no. 5, pp. 371-6, 2008, doi: 10.1080/13651820802291233.
[4] S. Paiella et al., "Radiofrequency ablation for locally advanced pancreatic cancer:
SMAD4 analysis segregates a responsive subgroup of patients," Langenbecks
Arch Surg, vol. 403, no. 2, pp. 213-220, Mar 2018, doi: 10.1007/s00423-017-
1627-0.
[5] H. Y. Ge et al., "High-intensity focused ultrasound treatment of late-stage
pancreatic body carcinoma: optimal tumor depth for safe ablation," Ultrasound
Med Biol, vol. 40, no. 5, pp. 947-55, May 2014, doi:
10.1016/j.ultrasmedbio.2013.11.020.
[6] Y. Keisari, "Tumor abolition and antitumor immunostimulation by physico-
chemical tumor ablation," Front Biosci (Landmark Ed), vol. 22, pp. 310-47, 2017.
234
[7] A. Sofuni et al., "Safety trial of high-intensity focused ultrasound therapy for
pancreatic cancer," World J Gastroenterol, vol. 20, no. 28, pp. 9570-7, Jul 28
2014, doi: 10.3748/wjg.v20.i28.9570.
[8] K. Sugimoto et al., "Irreversible Electroporation for Nonthermal Tumor Ablation in
Patients with Locally Advanced Pancreatic Cancer: Initial Clinical Experience in
Japan," Intern Med, vol. 57, no. 22, pp. 3225-3231, Nov 15 2018, doi:
10.2169/internalmedicine.0861-18.
[9] A. M. Ierardi et al., "Percutaneous ablation therapies of inoperable pancreatic
cancer: a systematic review," Ann Gastroenterol, vol. 28, no. 4, pp. 431-9, Oct-
Dec 2015. [Online]. Available: https://www.ncbi.nlm.nih.gov/pubmed/26424487.
[10] M. D'Onofrio et al., "Percutaneous ablation of pancreatic cancer," World J
Gastroenterol, vol. 22, no. 44, pp. 9661-9673, Nov 28 2016, doi:
10.3748/wjg.v22.i44.9661.
[11] M. Linecker, T. Pfammatter, P. Kambakamba, and M. L. DeOliveira, "Ablation
Strategies for Locally Advanced Pancreatic Cancer," Dig Surg, vol. 33, no. 4, pp.
351-9, 2016, doi: 10.1159/000445021.
[12] J. E. Parsons, C. A. Cain, G. D. Abrams, and J. B. Fowlkes, "Pulsed cavitational
ultrasound therapy for controlled tissue homogenization," Ultrasound Med Biol,
vol. 32, no. 1, pp. 115-29, Jan 2006, doi: 10.1016/j.ultrasmedbio.2005.09.005.
[13] E. Vlaisavljevich, A. Maxwell, L. Mancia, E. Johnsen, C. Cain, and Z. Xu,
"Visualizing the Histotripsy Process: Bubble Cloud-Cancer Cell Interactions in a
Tissue-Mimicking Environment," Ultrasound Med Biol, vol. 42, no. 10, pp. 2466-
77, Oct 2016, doi: 10.1016/j.ultrasmedbio.2016.05.018.
235
[14] Z. Xu, M. Raghavan, T. L. Hall, M. A. Mycek, and J. B. Fowlkes, "Evolution of
bubble clouds induced by pulsed cavitational ultrasound therapy - histotripsy,"
IEEE Trans Ultrason Ferroelectr Freq Control, vol. 55, no. 5, pp. 1122-32, May
2008, doi: 10.1109/TUFFC.2008.764.
[15] E. Vlaisavljevich et al., "Non-Invasive Liver Ablation Using Histotripsy: Preclinical
Safety Study in an In Vivo Porcine Model," Ultrasound Med Biol, vol. 43, no. 6,
pp. 1237-1251, Jun 2017, doi: 10.1016/j.ultrasmedbio.2017.01.016.
[16] Y. Kim, E. Vlaisavljevich, G. E. Owens, S. P. Allen, C. A. Cain, and Z. Xu, "In vivo
transcostal histotripsy therapy without aberration correction," Phys Med Biol, vol.
59, no. 11, pp. 2553-68, Jun 7 2014, doi: 10.1088/0031-9155/59/11/2553.
[17] E. Vlaisavljevich, Y. Kim, G. Owens, W. Roberts, C. Cain, and Z. Xu, "Effects of
tissue mechanical properties on susceptibility to histotripsy-induced tissue
damage," Phys Med Biol, vol. 59, no. 2, pp. 253-70, Jan 20 2014, doi:
10.1088/0031-9155/59/2/253.
[18] A. M. Lake, Z. Xu, J. E. Wilkinson, C. A. Cain, and W. W. Roberts, "Renal
ablation by histotripsy--does it spare the collecting system?," J Urol, vol. 179, no.
3, pp. 1150-4, Mar 2008, doi: 10.1016/j.juro.2007.10.033.
[19] T. Y. Wang, T. L. Hall, Z. Xu, J. B. Fowlkes, and C. A. Cain, "Imaging feedback of
histotripsy treatments using ultrasound shear wave elastography," IEEE Trans
Ultrason Ferroelectr Freq Control, vol. 59, no. 6, pp. 1167-81, Jun 2012, doi:
http://dx.doi.org/10.1109/TUFFC.2012.2307
10.1109/tuffc.2012.2307.
236
[20] J. E. Lundt, S. P. Allen, J. Shi, T. L. Hall, C. A. Cain, and Z. Xu, "Non-invasive,
Rapid Ablation of Tissue Volume Using Histotripsy," Ultrasound Med Biol, vol. 43,
no. 12, pp. 2834-2847, Dec 2017, doi: 10.1016/j.ultrasmedbio.2017.08.006.
[21] E. Vlaisavljevich et al., "Image-guided non-invasive ultrasound liver ablation
using histotripsy: feasibility study in an in vivo porcine model," Ultrasound Med
Biol, vol. 39, no. 8, pp. 1398-409, Aug 2013, doi:
10.1016/j.ultrasmedbio.2013.02.005.
[22] J. Vidal-Jove, E. Perich, and M. A. Del Castillo, "Ultrasound Guided High
Intensity Focused Ultrasound for malignant tumors: The Spanish experience of
survival advantage in stage III and IV pancreatic cancer," Ultrason Sonochem,
vol. 27, pp. 703-706, Nov 2015, doi: 10.1016/j.ultsonch.2015.05.026.
[23] J. E. Parsons, C. A. Cain, G. D. Abrams, and J. B. Fowlkes, "Pulsed cavitational
ultrasound therapy for controlled tissue homogenization," (in eng), Ultrasound in
Medicine and Biology, vol. 32, no. 1, pp. 115-29, Jan 2006, doi: S0301-
5629(05)00374-1 [pii]
10.1016/j.ultrasmedbio.2005.09.005.
[24] J. E. Parsons, C. A. Cain, and J. B. Fowlkes, "Cost-effective assembly of a basic
fiber-optic hydrophone for measurement of high-amplitude therapeutic ultrasound
fields," J Acoust Soc Am, vol. 119, no. 3, pp. 1432-40, Mar 2006, doi:
10.1121/1.2166708.
[25] H. R. Holmes et al., "Focused ultrasound extraction (FUSE) for the rapid
extraction of DNA from tissue matrices," (in English), Methods Ecol Evol, Oct 20
2020, doi: 10.1111/2041-210x.13505.
237
[26] J. Khirallah et al., "Nanoparticle-mediated histotripsy (NMH) using
perfluorohexane 'nanocones'," Phys Med Biol, vol. 64, no. 12, p. 125018, Jun 20
2019, doi: 10.1088/1361-6560/ab207e.
[27] K. C. Longo et al., "Robotically Assisted Sonic Therapy (RAST) for Noninvasive
Hepatic Ablation in a Porcine Model: Mitigation of Body Wall Damage with a
Modified Pulse Sequence," Cardiovasc Intervent Radiol, vol. 42, no. 7, pp. 1016-
1023, Jul 2019, doi: 10.1007/s00270-019-02215-8.
[28] A. R. Smolock et al., "Robotically Assisted Sonic Therapy as a Noninvasive
Nonthermal Ablation Modality: Proof of Concept in a Porcine Liver Model,"
Radiology, vol. 287, no. 2, pp. 485-493, May 2018, doi:
10.1148/radiol.2018171544.
[29] W. W. Roberts, D. Teofilovic, R. C. Jahnke, J. Patri, J. M. Risdahl, and J. A.
Bertolina, "Histotripsy of the prostate using a commercial system in a canine
model," The Journal of urology, vol. 191, no. 3, pp. 860-865, 2014.
[30] M. Shpoliansky, Y. Nevo, Y. Klein, and M. Amitai, "The Accuracy of Early Post-
Operative Abdominal CT Scan," Global Journal of Surgery, vol. 7, no. 1, pp. 19-
25, 2019.
[31] L. C. Sebeke et al., "Feasibility study of MR-guided pancreas ablation using high-
intensity focused ultrasound in a healthy swine model," International Journal of
Hyperthermia, vol. 37, no. 1, pp. 786-798, 2020.
[32] L. Ke et al., "The effect of intra-abdominal hypertension incorporating severe
acute pancreatitis in a porcine model," PloS one, vol. 7, no. 3, p. e33125, 2012.
238
[33] J. Otto, M. Afify, U. Jautz, V. Schumpelick, R. Tolba, and A. Schachtrupp,
"Histomorphologic and ultrastructural lesions of the pancreas in a porcine model
of intra-abdominal hypertension," Shock, vol. 33, no. 6, pp. 639-645, 2010.
[34] P. Marchetti, E. H. Finke, A. Gerasimidi-Vazeou, L. Falqui, D. W. Scharp, and P.
E. Lacy, "Automated large-scale isolation, in vitro function and
xenotransplantation of porcine islets of Langerhans," Transplantation, vol. 52, no.
2, pp. 209-213, 1991.
[35] M. Bower, L. Sherwood, Y. Li, and R. Martin, "Irreversible electroporation of the
pancreas: definitive local therapy without systemic effects," Journal of surgical
oncology, vol. 104, no. 1, pp. 22-28, 2011.
[36] C. B. Nahm, S. J. Connor, J. S. Samra, and A. Mittal, "Postoperative pancreatic
fistula: a review of traditional and emerging concepts," Clinical and experimental
gastroenterology, vol. 11, p. 105, 2018.
[37] C. Bassi et al., "The 2016 update of the International Study Group (ISGPS)
definition and grading of postoperative pancreatic fistula: 11 Years After," (in
eng), Surgery, vol. 161, no. 3, pp. 584-591, Mar 2017, doi:
10.1016/j.surg.2016.11.014.
239
Chapter 7. Immunological Effects of Histotripsy for Cancer Therapy
Alissa Hendricks-Wenger, Ruby Hutchison, Eli Vlaisavljevich, Irving C Allen
This chapter is excerpted from a manuscript in preparation for submission to Frontiers
of Oncology.
240
7.1 Abstract
Cancer is the second leading cause of death worldwide despite major advancements in
diagnosis and therapy over the past century. One of the most debilitating aspects of
cancer is the burden brought on by metastatic disease. Therefore, an ideal treatment
protocol would address not only debulking larger primary tumors but also circulating
tumor cells and distant metastases. To address this need, the use of immune
modulating therapies has become a pillar in the oncology armamentarium. A therapeutic
option that has recently emerged is the use of focal ablation therapies that can destroy
a tumor through various physical or mechanical mechanisms and release a cellular
lysate with the potential to stimulate an immune response. Histotripsy is a non-invasive,
non-ionizing, non-thermal, ultrasound guided ablation technology that has shown
promise over the past decade as a debulking therapy. As histotripsy therapies have
developed, the full picture of the accompanying immune response has revealed a wide
range of immunogenic mechanisms that include DAMP and anti-tumor mediator
release, changes in local cellular immune populations, development of a systemic
immune response, and therapeutic synergism with the inclusion of checkpoint inhibitor
therapies. These studies also suggest that there is a reproducible immune effect from
histotripsy therapies across multiple tumor types. Overall, the effects of histotripsy on
tumors show a positive effect on immunomodulation and clinical outcomes that could be
harnessed to provide systemic benefits to patients.
241
7.2 Keywords:
Abscopal Effect, Histotripsy, Focused Ultrasound, Immunomodulation
242
7.3 Introduction
Although cancer has plagued mankind throughout history, there still remains a
pressing need to improve treatment options. The core pillars of cancer therapy are
chemotherapy, surgery, radiation, immunotherapy, and ablation. The most common
treatment option for solid tumors remains surgical resection, even though many patients
are not candidates due to tumor size, location, or disease progression (Befeler and Di
Bisceglie, 2002; Chiorean and Coveler, 2015; Hsieh et al., 2017; Armengol et al., 2018).
Tumor ablation modalities have been developed as minimally and non-invasive
adjuvants or alternatives to surgery. These procedures include radiofrequency ablation
(RFA), microwave ablation, cryoablation, irreversible electroporation (IRE), high-
intensity focused ultrasound (HIFU), and histotripsy. While these procedures address
many of the issues with traditional surgery, there are still many areas where these
modalities can be improved.
While the goal of the field of oncology research is to find curative therapies, novel
cancer therapies frequently aim for non-curative endpoints such as increased
progression free survival, increased overall survival, decreased tumor burden, and
improved quality of life. In many cases, the therapeutic goals of local ablation therapy
also include a decrease in tumor burden, enhancement of drug delivery to tumors, and
the modulation of the immune system. While immune system modulation offers the
most promise in terms of long-term patient benefit, it is currently the least predictable
and understood benefit of local tumor ablation therapy. The most observable and
243
reliable benefit of local tumor ablation modalities is currently considered to be the
decrease in targeted tumor size (Adam et al., 2020). This can lead to improved patient
outcomes by debulking tumors, which can reduce the patient’s overall tumor burden,
improve local vasculature compression, and reduce tumor nutrient utilization.
7.4 The Role of Ablation in Immunomodulation
Given that one of the most debilitating aspects of cancer is the burden brought on
by metastatic disease, an ideal treatment protocol would address not only debulking
larger or debilitating tumors, but also circulating tumor cells and distant metastases.
Once cancer has spread from its primary site, focal debulking of individual tumors are
less likely to be capable of being a curative treatment option. Therefore, an ideal
debulking therapy would not only reduce the size, or fully eliminate, targeted tumors, but
would also stimulate the immune system to seek and destroy metastatic lesions
throughout the body. This phenomenon has been described as the abscopal effect
(Fig.1). There is increasing evidence that immune system activation and systemic
tumor elimination is critically important for the elimination of micrometastatic lesions
distal to the locally treated tumor, that may not be detectable at the time of treatment or
surgery. Before the abscopal effect was formally defined, it was first noted by physicians
at the turn of the 20th century who reported that local, targeted radiation of the spleen in
leukemia patients would cause changes in the patient’s bone marrow leading to the
remission of disease (Senn, 1903; Capps and Smith, 1907). These clinical findings in
combination with many mouse studies led to the abscopal effect being defined by the
244
early 1950’s to encompass the effects to distant organs or cancers after applying
ionizing radiation (Gunz, 1953). This effect was originally thought to only occur for
leukemia, where the radiation of the spleen would reduce or eliminate cancerous cells in
the body (Gunz, 1953; Hotchkiss and Block, 1962). However, over the subsequent
decades, additional reports of solid tumors displaying this effect periodically emerged.
For instance, the reduction of the sizes of distant metastases were reported after the
radiation of a single giant follicular lymphoma or a non-Hodgkin's lymphoma tumor
(Nobler, 1969; Antoniades et al., 1977). However, due to the scarcity of reports, the
abscopal effect in response to ionizing radiation of solid tumors, including renal cell
carcinoma, mammary carcinoma, and neuroblastoma, was frequently dismissed and
remained controversial for decades (Yang et al., 1992; Sanchez-Ortiz et al., 2003;
Demaria et al., 2004; Kaminski et al., 2005). To this day, the abscopal effect associated
with radiation therapy remains controversial (Wang et al., 2020). More recently, the
development of non-ionizing ablation therapies has led to an increase in the number of
abscopal effects reported. Thermal ablations, the first ablation therapies that were
developed and widely accepted, also have reported sporadic cases of a systemic
immune mediated effect where untreated, distal metastases were found to
spontaneously regress (Sanchez-Ortiz et al., 2003; Kondo et al., 2006; Kim et al.,
2008). One of these earlier case studies showed that the treatment of a primary renal
cell carcinoma tumor with RFA caused a spontaneous regression of pulmonary
metastases (Sanchez-Ortiz et al., 2003). On the other hand, as more non-thermal
ablation modalities are being developed pre-clinical studies have had more promising
correlations between focal ablation therapy and the systemic effects (Ringel-Scaia et
245
al., 2019; Scheffer et al., 2019; Qu et al., 2020). In general, even though there have
been promising trends that these therapies can offer systemic therapeutic effects for
patients, it should be noted that the reproducibility between patients, tumor types, and
therapy modalities is less than optimal (Kepp et al., 2019).
To achieve an abscopal effect, it is not sufficient to simply release non-specific
markers of tissue damage or cell death, if that were the case surgical resections would
stimulate systemic effects. Instead, there is a need to release specific tumor associated
antigens that allow the immune system to build a targeted response (Fig.1). When
focused, local ablation therapies break up the stromal tissues of a tumor this can create
an increased opportunity for immune cells to physically access the tumor (Dromi et al.,
2009; Lu et al., 2009). As the cancer cells are ablated, there is an increased abundance
of accessible tumor antigens that can be recognized by infiltrating antigen presenting
Figure 7-1 Focal Treatment of Targeted Tumor and Mechanism for Systemic Tumor Control. The right diagram demonstrates the principal of the abscopal effect, where the treatment of one tumor can cause other tumors in the body to shrink or be eliminated to varying degrees due to immune system engagement. The left diagram depicts the simplified mechanism for achieving a systemic effect from focal therapies. DC: Dendritic cell. MHC: Major Histocompatibility Complex. TCR: T cell receptor.
246
cells (APC), such as dendritic cells (den Brok et al., 2004; Kim et al., 2008). These
APCs can in turn activate lymphocytes to guide a more precise immune response.
Locally, activation of cytotoxic CD8+ T cells can increase the clearance of any surviving
cancer cells within the area treated with the tumor ablation modality (Ito et al., 2019).
For the strongest systemic immune effects, after ablation, dendritic cells will process the
available tumor associated antigens and migrate to the nearest draining lymph nodes
and the spleen. Once in the lymphoid tissues, the dendritic cells present the antigens
through MHC class I or class II receptors to naïve CD4+ and CD8+ T cells, respectively,
along with co-stimulation of CD28 with CD80/86. Studies that show an increase in tumor
specific lymphocytes in secondary lymphoid organs or in peripheral blood have also
established the correlation between these activated cells and systemic decreases in
tumor burden (Wissniowski et al., 2003; Wu et al., 2004; Widenmeyer et al., 2011;
Leuchte et al., 2020). More recently, additional studies have also defined B cell
activation, the maintenance of plasma cells, and even the generation of tumor specific
antibodies following local tumor ablation and dendritic cell activation (Widenmeyer et al.,
2011; Leuchte et al., 2020). While the activation of T and B lymphocytes is a core
feature of the systemic effects of local tumor ablation therapy, there are many
inconsistencies. For example, a study treating hepatocellular carcinoma with RFA
reported that only 91 of the 178 patients that had an elevated neutrophil-to-lymphocyte
ratio, which is an often-used biomarker of systemic immune system activation,
independently correlated to a significantly increased survival when compared to patients
that did not see the increased ratio (Dan et al., 2013). Similarly, in a prospective study
investigating the long-term effects of RFA after treating secondary liver tumors, it was
247
found that only 6 of 49 patients studied had increased antibodies, CD4+ T cells, and/or
CD8+ T cells months after treatment (Widenmeyer et al., 2011). While the exact
mechanisms underlying the inconsistent reports of the abscopal effect and systemic
anti-tumor immune responses are yet to be fully defined, changes in the local tumor
microenvironment have been postulated to significantly impact treatment success.
7.4.1 The Tumor Microenvironment and Tumor Associated Immune Cells
To determine the cause of systemic anti-tumor immunity, and therefore how to
modulate it, there is a need to understand what is occurring within the tumor
microenvironment. Within the heterogeneous cell populations of most tumors, there are
a variety of tumor-associated immune cells. In an ideal case, these leukocytes are
eliminating malignant cells. However, in the case of many cancers, the leukocytes are
reprogrammed, or polarized, to support tumor escape and function to promote
tumorigenesis through the generation of a relatively immunosuppressive tumor
microenvironment. These tumors are often referred to as being immunologically “cold”
due to their lack of inflammation and active immune suppression. With multiple types of
pro-tumor immune cells that produce factors that can improve the tumor niche, it is
difficult for the pro-inflammatory/anti-tumor immune response to overcome this “cold”
environment.
The leukocytes most commonly associated with these anti-inflammatory and tumor
promoting properties include tumor-associated macrophages (TAM), myeloid-derived
suppressor cells (MDSC), and tumor-associated neutrophils (TAN) (Fig.2). TAMs are
248
generally defined as CD45+ Ly6C- MHCII+ CD11b+ (Ostuni et al., 2015; Fu et al.,
2020). TAMs are also broken down into subsets based on polarization to the M1
macrophage-like phenotype (pro-inflammatory/anti-tumor) CD68+ CD80/86+ CD11c+
iNOS+ and the M2 macrophage-like phenotype (pro-tumor) CD163+ CD204+ CD206+
Arg1+, with M2-like TAMs being the most immunosuppressive (Wu et al., 2020; Zhou et
al., 2020). It should be noted that while the M1-like and M2-like TAMs do share similar
expression patterns and activities, they are not the same as the traditional M1 and M2
polarized macrophages. MDSCs are generally defined as CD45+ CD11b+ CD11c-
MHCII- as well as CD15+ and CD66b+ in humans (Ko et al., 2009; Rodriguez et al.,
2009; Youn and Gabrilovich, 2010). MDSCs are broken into two main subtypes:
monocytic MDSCs Ly6C+ Ly6G- and granulocytic MDSCs Ly6C- Ly6G+ (Youn and
Gabrilovich, 2010). When generally searching for MDSCs, the GR-1 antibody has been
used given that it binds to both Ly6C and Ly6G (Youn and Gabrilovich, 2010). TANs are
defined as CD45+ CD11b+ Ly6G+ and have been reported to express CD66 and CD15
in human cancers (Fridlender et al., 2009; Hajizadeh et al., 2020; Wu and Zhang, 2020).
It should be noted that the markers that have been established for TANs are
indistinguishable from traditional neutrophils, with the most established phenotypic
difference between the two cell groups being that neutrophils have a half-life of 6-8
hours whereas TANs have a significantly increased lifespan (an additional 12-24 hours)
caused by an inhibition of apoptotic pathways and support from cytokines within the
tumor microenvironment (Wislez et al., 2001; van Raam et al., 2008; Summers et al.,
2010; Trellakis et al., 2011; Hajizadeh et al., 2020). Similar in nomenclature to TAMs,
TANs can also be divided into N1 pro-inflammatory and N2 pro-tumor subtypes,
249
however due to limitations in markers there is not an established panel for differentiating
N1 from N2 TANs (Fridlender et al., 2009).
Figure 7-2 Pro-Tumor Immune Cells. CD66, CD66b, and CD15 are human specific markers. Gr-1 represents both Ly6C and Ly6G, therefore does not distinguish the MDSC subtypes, MDSCs are monocytic when Ly6C+ and granulocytic when Ly6G+. Most of the markers available for pro-tumor immune cells are surface receptors and ligands, exceptions listed here include the intracellular protein Arg1 and the transcription factor FOXP3.
250
These anti-inflammatory leukocytes are typically distributed throughout the tumor
and margins (Fig.2) and function beyond simply being tumor associated. These
leukocytes are frequently referred to as being pro-tumor given that the range of their
activities can actively promote tumor growth and development. These functions include
reducing the impact of anti-tumor immune responses, recruiting additional tumor
supporting cells, helping the tumor grow by stimulating angiogenesis and secreting
growth factors, and assisting in the epithelial-to-mesenchymal transition (Najafi et al.,
2019; Tesi, 2019; Wu et al., 2019). For TAMs, , this includes producing anti-
inflammatory mediators such as IL-10, Arg1, and TGF-β (Lang et al., 2002; Pang et al.,
2017; Arlauckas et al., 2018), promoting the expression of checkpoint inhibitors to
suppress T cells (Pang et al., 2017; Arlauckas et al., 2018), inducing cancer stem cell
proliferation via IL-6 signaling through STAT3 (Wan et al., 2014), and producing VEGF
to stimulate angiogenesis (Bolat et al., 2006; Hu et al., 2017). These effects are also
seen in TANs and MDSCs, which have both been shown to produce similar profiles of
cytokines, checkpoint inhibitors, and growth factors (Gao et al., 2020; Hajizadeh et al.,
2020; Wu and Zhang, 2020; Mabuchi et al., 2021; Zhou et al., 2021).
In addition to these monocyte and granulocyte derived cells, lymphocytes can also
aid in maintaining this “cold” tumor microenvironment. Regulatory T cells, defined as
CD3+ CD4+ CD25+ FOXP3+, can suppress the adaptive immune response (Tanaka
and Sakaguchi, 2019). While MDSCs, TANs, and TAMs are commonly identified as
tumor promoting cells with specific markers and functions, the role of T regulatory cells
is more situationally specific and their presence and ratio are not necessarily a sign of a
251
“cold” tumor microenvironment. In most cases, the presence of a high ratio of T
regulatory cells, known for inhibiting the function of other T cells, is a sign of a “cold”
tumor microenvironment. Clinically, in ovarian carcinoma it has been established that
the recruitment of T regulatory cells into the tumor stroma leads to decreased survival of
patients (Curiel et al., 2004). Additionally, for pancreatic cancer, the T regulatory cells
have been found to aid in the pro-carcinogenic inflammation driven by T helper 17 cells
(Vizio et al., 2012). These poor prognostic correlations in patients is tied to the fact that
T regulatory cells can suppress the function of T helper cells through the production of
immunosuppressive cytokines, including IL-10 and TGF-β (Levings et al., 2002;
Kindlund et al., 2017), secrete perforins and granzymes to directly destroy effector T
cells and B cells (Cao et al., 2007), and express the checkpoint inhibitor CTLA-4 to
suppress APC activity (Kalathil et al., 2013). There have also been reports of T
regulatory cells directly increasing the growth of tumors through the stimulation of
cancer stem cells via the NF-κB-IL6-STAT4 signaling axis (Liu et al., 2021).
On the other hand, there are reports of certain cancers, such as colorectal cancer,
which have worse prognoses for patients that have a reduced presence of T regulatory
cells within their tumors (Salama et al., 2009; Ladoire et al., 2011). A proposed
mechanism for the seemingly contradictory immune promoting function of T regulatory
cells in colorectal cancer is associated with the location of these tumors. In the
gastrointestinal tract, T regulatory cells are primed against the microbiota of the
intestinal space instead of the cancer (Ladoire et al., 2011). This secondary, non-
tumoral target for the immune system allows for the immune response to become more
252
robust in the colon, which can in turn destroy the cancerous cells, without being
hampered by the anti-inflammatory control of the tumor microenvironment.
7.4.2 Mechanisms for Shifting the Tumor Microenvironment to Pro-Inflammatory
The largest difference between a “cold” and a “hot” tumor microenvironment is
typically considered to be related to the infiltration of anti-tumor, pro-inflammatory T
cells in “hot” or inflamed tumors (Gajewski, 2015). Checkpoint inhibitors are emerging
as the primary therapeutics used to facilitate the shift in the tumor microenvironment
from “cold” to “hot” or immunologically active (Wilky, 2019). In the case of many
cancers, the checkpoint inhibiting pathways are hijacked to prevent immune cells from
eliminating unhealthy or cancerous cells. The two most heavily studied checkpoint
inhibitor pathways in the tumor ablation field are CTLA-4 to CD80/86, blocking the
normal co-stimulatory role of CD80/86 in T cell activation, and PD-1 to PDL1/2 (Fig.3).
CTLA-4 is a checkpoint inhibitor most commonly associated with antigen presentation in
the lymph nodes that minimizes the proliferation of T cells (Wilky, 2019).
Therapeutically, when CTLA-4 on T cells is blocked with a targeted antibody, there is an
increase in overall T cell proliferation and an increased probability of forming a tumor-
specific immune response. While anti-CTLA-4 is used to increase T cell proliferation,
PD-1 inhibitors decrease T cell inhibition and improve function. Under normal
conditions, PD-1 inhibits the activity of T cells and prevents the targeting of healthy host
cells by T lymphocytes (Wilky, 2019). This pathway in healthy tissues prevents
overzealous immune responses that can drive autoimmunity. However, some tumor
253
cells hijack this mechanism and increase the expression of the PD-1 ligands (PD-L1/2),
which artificially reduce the activity of anti-tumor T cells within the tumor
microenvironment. The use of antibodies targeting the PD-1 pathway for cancer therapy
has shown increased levels of infiltrating T cells in tumors that express the PD-L1/2
surface receptors. While PD-1 pathway inhibition has a more tumor-specific therapeutic
effect, and typically fewer side effects compared to CTLA-4 based therapies, there is
still a need to improve the implementation of checkpoint inhibition strategies. This is
especially true in immunologically “cold” tumors where these checkpoint targeted
therapeutics have proven to be minimally effective. Unfortunately, even in cases where
these therapies work there are often significant side effects. For example, the systemic
increase in T cell abundance often leads to autoimmune disease-like symptoms such as
fatigue, nausea, vomiting, diarrhea, arthritis, dermatitis, and myalgia (Arnaud‐Coffin et
al., 2019).
Figure 7-3 The Role of Check Point Inhibitors. Antigen Presenting Cells (APC).
254
Beyond the biochemical changes in the tumor microenvironment that work to subvert
immune system detection and elimination, physical barriers that can have significant
detrimental effects on therapeutic approaches are also present. Indeed, many tumors
adopt organ-like structures and extensive fibrosis that build highly complex physical
barriers (Plate, 2011; Jiang et al., 2017). Given that the checkpoint inhibitor’s
mechanism is to improve immune system activation and targeted killing of tumor cells, it
is critical that leukocytes have physical access to the malignant cells. In an attempt aid
the checkpoint inhibitor therapies, ablation modalities have been extensively studied in
an attempt to better understand their ability to modulate the immunological aspects of
the tumor microenvironment. While debulking the tumor and reducing a patient’s overall
tumor burden is the primary goal of focal tumor ablation therapies, improved immune
system access is proving to be an added benefit that can shift the local tumor
microenvironment from “cold” to “hot.”
Ablation therapies with a thermal effect have sporadically seen pro-inflammatory
changes in the tumor microenvironment in the weeks and months following treatment
(Saccomandi et al., 2018; Yin et al., 2018; Minami et al., 2019; Xie et al., 2019).
However, in terms of immunomodulatory success, non-thermal modalities seem to have
more reproducibility, predictability, and improved pre-clinical and clinical success
(Bastianpillai et al., 2015; Beitel-White et al., 2019; Pandit et al., 2019; Ringel-Scaia et
al., 2019). The leading hypothesis regarding differences between modalities theorizes
that while both thermal and non-thermal ablation approaches kill tumor cells, non-
thermal ablation modalities release higher levels of native tumor-specific antigens that
255
have not been distorted or denatured by heat. For example, one in vitro study showed
that intact neoantigens are released in significantly higher magnitudes from cells treated
with cryoablation and IRE compared to thermal ablation. Not only were there more
tumor antigens released, these antigens were also more potent at stimulating dendritic
cell antigen presentation and cytotoxic CD8+ T cell activation (Shao et al., 2019).
Activated lymphocytes, either in response to tumor-specific antigens or more generally
in response to innate immune system activation driven by the increase in damage-
associated molecular patterns (DAMPs) further promote the shift from a “cold” to a “hot”
tumor microenvironment (Chandra et al., 2016; Domingo-Musibay et al., 2017; Ringel-
Scaia et al., 2019). Thus, it is becoming clear that non-thermal ablation modalities are
capable of inducing robust local and systemic anti-tumor responses, as either an
independent or an adjuvant therapy with immunotherapeutics, to offer significant
promise beyond just focal tumor debulking.
7.5 Histotripsy: Non-Thermal, Non-Invasive Tumor Ablation
Histotripsy is a non-thermal focused ultrasound therapy that utilizes microsecond
(cavitation-cloud histotripsy, CCH) or millisecond (boiling histotripsy, BH) pulsing
regimens to generate cavitation bubble clouds that leads to precise non-thermal tumor
ablation (Fig.4) (Bader et al., 2019). In addition to histotripsy, other non-thermal focused
ultrasound methods that induce cavitation and do not induce substantial thermal effects
are broadly referred to as mechanical HIFU (mHIFU), as opposed to conventional HIFU
that typically refers to thermal ablation. For the purposes of this study, thermal HIFU will
256
be referred to as tHIFU to delineate from mHIFU procedures, which will be the focus of
this review.
In histotripsy, the rapid expansion and collapse of cavitation bubbles ablate tissues
into acellular debris (Vlaisavljevich et al., 2016a; Vlaisavljevich et al., 2017; Smolock et
al., 2018). This non-thermal ablation process is well-established and has resulted in
important hallmarks of histotripsy, including high precision and tissue selectivity
(Vlaisavljevich et al., 2013a; Lin et al., 2014; Vlaisavljevich et al., 2015a). For example,
tissues with a higher Young’s Modulus, such as vasculature and collecting ducts, are
more resistant to damage (Lake et al., 2008; Hall et al., 2009; Hempel et al., 2011;
Vlaisavljevich et al., 2013b; Vlaisavljevich et al., 2014a; Vlaisavljevich et al., 2014b;
Khokhlova et al., 2015; Vlaisavljevich et al., 2015b). Unlike the vast majority of other
ablation therapies, because histotripsy is non-thermal, it is not affected by the heat-sink
Figure 7-4 Histotripsy Schematic. Therapeutic ultrasound transducer positioned outside of the body, focuses ultrasound waves within a targeted tissue generating a cavitation bubble cloud.
257
effect, and therefore remains safe and efficacious for use near the vasculature (Thanos
et al., 2008; Pillai et al., 2015). Additional benefits of histotripsy include real-time
imaging feedback with standard imaging (ultrasound, MRI, CT), highly precise-
millimeter accuracy, and the ability to treat tumors of arbitrary sizes and shapes (Hall et
al., 2009; Smolock AR, 2018). After treatment, tissues treated with histotripsy have
shown more rapid dissolution of the ablated tissues compared to other ablation
modalities (Vlaisavljevich et al., 2016a; Smolock AR, 2018). For instance, CCH ablation
of healthy rat livers showed rapid involution of treated volumes, granulation, and growth
of healthy hepatocytes within 28 days with minimal scarring (Vlaisavljevich et al.,
2016a). This healing process is more rapid than what has been reported for other
ablation modalities, such as RFA in humans, which has been reported as causing
“thermal fixation,” where the necrotic mass is still present 14 months post-treatment due
to thermal denaturation of the tissue leaving it resistant to being broken down by normal
pathways (Coad et al., 2003).
In mouse models, histotripsy ablation of hepatic, renal, neuroblastoma, and
melanoma tumors has resulted in significantly increased survival (Schade, 2018;
Worlikar et al., 2019; Eranki et al., 2020; Qu et al., 2020). In addition to debulking
tumors, recent evidence has suggested that histotripsy has the capability to induce a
systemic immune response, as evidenced by the attenuation of metastasis and an
improvement in local and distant disease with combination immunotherapy (Worlikar et
al., 2018; Schade et al., 2019; Qu et al., 2020). This effect has been shown in single
tumor treatments, and has also shown abscopal-like decreases in contralateral tumor
growth in a separate untreated tumor (Schade et al., 2019; Eranki et al., 2020; Qu et al.,
258
2020). This effect was found to be modest, but statistically significant with histotripsy
ablation compared to the insignificant trend in mice treated with irradiation and
radiofrequency ablation (Qu et al., 2020). The molecular immunological effects caused
by histotripsy, independently and compared to other ablation modalities, can best be
summarized by the proposed mechanism shown in Figure 5 that is described in the
sections below.
7.6 Immunologic Effects of Histotripsy
The immunological responses to histotripsy ablation through cavitation,
regardless of the sub-therapy being applied, are theoretically similar due to comparable
effects on targeted tissues. Cavitation breaks down tissues, ablating cells into
subcellular fragments and acellular debris (Vlaisavljevich et al., 2016b). To date, there
have been no studies to suggest that mHIFU, BH, nor CCH have any significant
differences in immunologic responses. Therefore, for this review, the immune
responses to these modalities will be reviewed together.
7.6.1 Decreases in Pro-Tumor Immune Cells
Being able to reduce the magnitude of tumor supporting immune cells present
within the tumor microenvironment is pertinent due to the correlation of these cells with
poorer outcomes (Zamarron and Chen, 2011; Fleming et al., 2018; Wu et al., 2019).
There have been multiple studies on thermal and non-thermal ablation modalities that
show either the therapy directly eliminates cells that support the tumor
259
microenvironment or the damage initiated by the ablation reprograms the leukocytes
within the tumor microenvironment to shift the microenvironment to one that is more
harsh towards tumor progression (Keisari, 2017; Kepp et al., 2019).
Figure 7-5 Schematic of Histotripsy’s Postulated Roles in Immune System Modulation. Immune processes, cells, and molecules that have been shown to be modulated by focused ultrasound therapies after mechanical ablation are summarized based upon immune system functions: pro-tumor immune cells, cellular immunity, and systemic immunity.
260
In the case of histotripsy, there has been one report from Phak et al. demonstrating the
supernatant from cells treated with BH in vitro polarizes naïve macrophages (M0) into
anti-tumor macrophages (M1) and the redifferentiation of tumor associated
macrophages (M2-like phenotype) to the more inflammatory M1 phenotype (Pahk et al.,
2019). In this study, breast cancer cells were treated with BH at varying doses in vitro
and the supernatant collected. Analysis found increased pro-inflammatory signaling
molecules including TNF, which is a potent and well established M1 stimulating cytokine
(Wang and Lin, 2008; Kratochvill et al., 2015). After THP-1 human monocyte cells were
cultured with the BH supernatant they displayed physical and morphological changes
consistent with polarization to M1 macrophages and showed significantly elevated gene
expression consistent with M1 polarization (Pahk et al., 2019). Together, the data
presented in this study suggest that monocytes and macrophages near the histotripsy
ablation region polarize to pro-inflammatory, anti-tumor phenotypes. Likewise, Phak et
al. continued in vitro studies showing that the BH supernatant can also stimulate M2
cells to repolarize to M1, thus leading to a decrease in M2 pro-tumor macrophages
(Fig.5). Thus, these data further suggest that any tumor-associated macrophages
remaining within the treatment zone have the potential to repolarize from an M2-like
state to M1, similar to the cells at the margins of treatment. If this can be recapitulated in
vivo, this would help further decrease the presence of pro-tumor immune cells and
could significantly alter the shift the tumor microenvironment from “cold” to “hot.”
While it might seem safe to assume that any pro-tumor cells within the tumor
microenvironment of treatment zone in vivo would be ablated due to the subcellular
fractionation of histotripsy, there is still a window for any cells that survive from a partial-
261
ablation to proliferate and maintain the pro-tumor tumor microenvironment. No literature
confirms this hypothesis for histotripsy ablation in vivo. However, these findings are
consistent with results following RFA, microwave ablation, and IRE, which can all
reduce the presence of tumor supporting immune cells in the tumor microenvironment
(Keisari, 2017; Dai et al., 2021). This dynamic has also been seen in human patients.
For instance, studies of pancreatic adenocarcinoma patients treated with IRE found a
decrease in T regulatory cells in peripheral blood in the days following treatment (Beitel-
White et al., 2019; Pandit et al., 2019). Another study of patients with primary liver
tumors treated with RFA showed a strong correlation between the decrease in MDSCs
and the increase in patients’ survival (Wang et al., 2016). The importance of the
attenuation of pro-tumor immune cells, their decrease in function, and reduction in
magnitude after ablation are all generally correlated with reduced survival in human
patients. Therefore, it is important for the field of histotripsy to further understand the
effects of the ablation on this aspect of the immunomodulatory mechanism.
7.6.2 Histotripsy Produces Damage Associated Molecular Patterns that Directly
Increase Local Cellular Immune Responses
Histotripsy generates subcellular fragments of targeted cells through mechanical
fractionation (Vlaisavljevich et al., 2016b). This, in turn, releases a multitude of damage
associated molecular patterns (DAMPs) that have the potential to stimulate the innate
immune system and significantly alter the tumor microenvironment. DAMPs are
molecules that are found extracellularly following most forms of pathological cell death.
262
These molecules are readily identified by the immune system, which in turn elicits a
robust innate immune response. Several DAMPs have been observed and evaluated in
the context of tumor ablation, including the endoplasmic reticulum associated protein
calreticulin (CRT), the non-histone nuclear binding protein HMGB1, the intracellular
energy molecule adenosine 5’-triphosphate (ATP), and various heat shock proteins
(HSP) that were originally identified as DAMPs released after exposure to elevated
temperatures (Harris and Andersson, 2004; Pisetsky, 2011; Korbelik et al., 2015; Chen
et al., 2019; Fodor et al., 2020). Once released from the injured cells or neighboring
cells following damage, these molecules can stimulate a robust immune response by
bindig to receptors known as pattern recognition receptors. As an example, HMGB1
commonly activates the Toll-like Receptors 2 and 4, which classically signals through
NF-κB to drive multiple biological responses, including inflammation and cell death (van
Beijnum et al., 2008).
These DAMPs are emerging as key mediators of the ensuing immune response
following histotripsy. Specifically, following histotripsy treatment of cancer spheroids,
HMGB1, CRT, and HSP were identified in cell supernatants and correlated to levels
expected from within the tumor after an in vivo ablation (Pahk et al., 2019). Additionally,
murine tumors treated with histotripsy have been found to have elevated levels of
HMGB1, CRT, and HSP within the tumor and increased HMGB1 has been routinely
found in serum following treatment (Hu et al., 2005; Schade, 2018; Qu et al., 2020).
DAMPs released in serum function to systemically prime the immune system and can
function to recruit increased circulating leukocytes that can ultimately congregate at the
site of tissue injury following ablation. Together, the local and systemic presence of
263
DAMPs aid additional immune cell migration, shift the tumor microenvironment, and
may eventually serve as effective biomarkers of ablation success.
While DAMP release has been confirmed following histotripsy, it is also critical to
determine if these signals are in their naïve and unaltered structural configurations. For
thermal ablation modalities, it has been well established that proteins can be denatured
under high heat settings, making them less efficacious and predictable in stimulating the
immune system. One study investigated the difference in the release of ATP and hsp60
between mHIFU and tHIFU (Hu et al., 2005). This study found that mHIFU is capable of
releasing higher levels of both molecules after treatment, and that the DAMPs released
from mHIFU were more capable of stimulating downstream immunologic changes, such
as activating dendritic cells (Hu et al., 2005). The ability of mHIFU to release a higher
magnitude of active DAMPs compared to tHIFU adds to the hypothesis that non-thermal
ablation therapies are more immunomodulatory compared to thermal modalities.
These data point to histotripsy therapies having the ability to release molecules that
can drive anti-tumor immune system activation, for example by initiating a robust innate
immune response. However, the data associated with the specific DAMPs released
following histotripsy is not consistent across studies. For example, a recent study of
subcutaneous murine neuroblastoma treated with mHIFU showed no significant
changes in HMGB1 levels (Eranki et al., 2020). This could be due to a difference in
tumor types. For example, neuroblastoma did not see the change in DAMPs after
histotripsy that was observed in colon adenocarcinoma, renal cell carcinoma, and
breast adenocarcinoma reported elsewhere (Hu et al., 2005; Schade, 2018; Eranki et
al., 2020; Qu et al., 2020). Alternatively, there are potential differences in histotripsy and
264
other mHIFU ablation modalities that are not fully understood and may have played a
role. Another possibility is that differences in DAMP release between a partial ablation
versus a more complete treatment are responsible for these differences. All of these
variables could potentially affect the quality and quantity of DAMPs released. Likewise,
the actual DAMPs generated could significantly vary depending on a number of
experimental factors. Although the aforementioned DAMPs are currently the most
defined following histotripsy, any DAMP could conceivably function as an activator of
the innate immune system after ablation. Future investigation into the effects of other
DAMPs, such as nuclear and mitochondrial DNA, reactive oxygen species, and calcium
ions could prove useful for improving adjuvant therapies (Gong et al., 2019). Additional
studies are needed to determine any potential difference in the quantity and quality of
the release of DAMPs from the different mechanical focused ultrasound therapies.
7.6.3 Pro-Inflammatory Cytokines and Chemokines Associated with Histotripsy Based
Tumor Ablation Modalities
Cytokines and chemokines are critical for cell-to-cell communication and immune
system modulation following damage (Fig. 5). These molecules stimulate the
differentiation and activation of local immune cells and systemically recruit additional
cells to the site of damage. One of the critical cytokines found to be significantly altered
in multiple histotripsy studies is INFγ (Pahk et al., 2019; Schade et al., 2019; Eranki et
al., 2020; Qu et al., 2020). In an in vitro study on breast adenocarcinoma and an in vivo
murine study of neuroblastoma, there was an approximately 2-fold increase in INFγ
265
after histotripsy (Pahk et al., 2019; Eranki et al., 2020). However, in an in vivo study
where Eker rat renal cell carcinoma tumors were treated with histotripsy, there was a
significant decrease, ~6pg/mL down to ~2pg/mL, in treated kidneys (Schade et al.,
2019). This decrease was attributed to the native kidney tissues’ defense mechanism
to protect against acute and chronic kidney injuries (Chung and Lan, 2011). The
importance of changes to INFγ is highlighted by its role in activating anti-tumor APCs
and effector cells, inducing ischemia within the tumor by acting on the endothelial cells
thus reducing the stability of intratumoral vasculature, and initiating tumor cell death
through the activation of apoptosis and necroptosis pathways (Ni and Lu, 2018).
However, IFNγ is also a double-edged sword in the tumor microenvironment. In addition
to its function as a potent pro-inflammatory mediator, IFNγ can also up-regulate PD-L1
in some tumors as part of the IRF1 axis, resulting in immune evasion (Mimura et al.,
2018).
While INFγ is the most consistently reported inflammatory mediator across
therapies and tumor types, other important cytokines have been reported as either
significantly or notably upregulated in the days after histotripsy treatment including IL-6,
IL-2, TNF, IL-8, IL-13, and IL-10 (Schade, 2018; Pahk et al., 2019; Eranki et al., 2020;
Qu et al., 2020). While more research is needed to more accurately determine the state
of cytokines in specific tumor types after mechanical ablation, the general picture
appears positive. However, high levels of IL-6, IL-10, and IL-8 in serum have been
reported to negatively correlate with survival in pancreatic cancer patients, due to their
role in stimulating inflammation and adverse changes in the tumor microenvironment
that spurs the growth and development of pancreatic tumors (Feng et al., 2018; Shadhu
266
and Xi, 2019). Although this could correlate to a foreboding warning against histotripsy
therapies, at least for pancreatic applications, the induction of these pathways actually
correlates with improved anti-tumor immune responses and increased survival in mice
(Qu et al., 2020). It is certainly possible that these results simply reflect the commonly
promoted differences between mice and humans. However, it is also possible that
additional mechanistic insight may be necessary to better translate the findings from
rodents to human patients.
In addition to the cytokines released there are changes in the serum levels of
multiple growth factors as well. For example, after mHIFU ablation of neuroblastomas,
both an increase in GM-CSF and a decrease in VEGF were found within 24 hours of
treatment (Eranki et al., 2020). Increased levels of GM-CSF are positively correlated
with increased APC cell differentiation and activation. Decreased levels of VEGF are
positively correlated with better clinical outcomes, due to the rate of vascularization of
tumors affecting tumor growth and immunosuppressive effects (Lapeyre-Prost et al.,
2017). The finding that histotripsy can increase GM-CSF and decrease VEGF adds to
the potential for histotripsy to shift the tumor promoting immune microenvironment to
one that is more proinflammatory and tumor suppressive.
267
7.6.4 Histotripsy Significantly Alters Immune Cell Populations Systemically and in the
Tumor Microenvironment
With the increased release of DAMPs and anti-tumor mediators, there is a
resultant change in intra-tumoral immune cell populations. In response to damage, the
cells associated with innate immunity are the most rapidly recruited. For histotripsy
ablation, this has been reported to include neutrophils, natural killer cells, dendritic cells,
and macrophages (Fig. 5). In an in vitro experiment, the supernatant of mHIFU treated
murine colon adenocarcinoma cells was shown to activate macrophages (Hu et al.,
2005). This macrophage activation was established through the release of TNF from the
macrophages, which was significantly greater after mHIFU compared to both tHIFU and
controls (Hu et al., 2005). Additionally, in vivo treatment of melanoma tumors with CCH
showed increases in neutrophils, natural killer cells, dendritic cells, and macrophages
10 days after treatment (Qu et al., 2020). This increase in intratumoral immune cell
populations indicate that the tumor microenvironment has become more
immunologically “hot” and these cell populations typically correlate to pro-inflammatory
and anti-tumor changes in the tumor microenvironment.
Beyond innate immune cells, modulation of adaptive immune cells after ablation
has also been strongly correlated with clinical success (Keisari, 2017; Kepp et al.,
2019). In the same melanoma CCH ablation study, increases in T helper cells and B
cells within the treated tumors were observed 10 days post-treatment (Qu et al., 2020).
In addition to the increase in these adaptive immune cell populations, the study also
found a decrease in intratumoral T regulatory cells (Qu et al., 2020). The increase in
268
helper T cells and B cells is a further indication that the TME has shifted to a more
permissible, pro-inflammatory environment and further illustrates the improved
accessibility of these cells to the tumor. In general, the influx of leukocytes appears to
reflect the early induction of the innate immune system and eventually yields to cells
associated with a robust adaptive immune response over the course of a few weeks
following treatment. Together, these changes are predicted to have significant impacts
on the local and systemic effects on metastatic lesions. However, it is critical to note
that, while the increased presence of these pro-inflammatory immune cells within the
residual treated tumor does hint at a stronger and more robust overall immune
response, these cells must be specifically primed for tumor antigens found and either in
the circulation or in secondary lymphoid organs to effectively control the overall disease
burden.
7.6.5 Improved Engagement of the Adaptive Immune System and Increases in the
Systemic Anti-Tumor Immune Response
The initiation of the systemic anti-tumor immune response requires the
generation of both high quantity and high-quality tumor antigens. Previous studies with
thermal ablation, cryoablation, and IRE established that the non-thermal ablation
modalities appear to release antigens that are significantly better at driving predictable
and effective antigen presentation (Shao et al., 2019). This ultimately results in
improved systemic immune responses (Shao et al., 2019). Although an explicit study
has yet to explore if histotripsy ablation negatively impacts the quality of released
269
antigens, studies have been conducted that indirectly evaluate this mechanism. A study
using the B16F10 murine melanoma cell line and a transgenic variant of the cell line
revealed an increase in the immunomodulatory effects of histotripsy in the tumors that
expressed an immunogenically active antigen (Qu et al., 2020). In this study, only mice
that had tumors transfected with the potential antigen GP33, a known antigen from
lymphocytic choriomeningitis virus, and were also treated with histotripsy, generated
CD8+ T cells that were capable of producing IFNγ after stimulation with IL-2 and
brefeldin A. These two agents stimulate memory CD8+ T cells. Together, these data
imply that transfected antigens are still viable after histotripsy treatment and are able to
stimulate an adaptive immune response.
The activation and migration of dendritic cells to tumor-draining lymph nodes and
the spleen are further evidence of systemic immune system activation. Early studies
into dendritic cells found the supernatants collected from in vitro mHIFU treatments
were able to stimulate immature dendritic cells to express higher levels of CD80 and
CD86, which are markers of activation, and enhanced secretion of IL-12, which are
utilized by dendritic cells to activate both CD8+ cytotoxic and CD4+ helper T cells
(Watford et al., 2003; Hu et al., 2005; Henry et al., 2008). Additional murine studies with
various tumor types further established the migration of, or the increase in, the number
of dendritic cells to the tumor-draining lymph nodes and spleen (Hu et al., 2007; Huang
et al., 2012; Eranki et al., 2020). The higher presence of this critical antigen-presenting
cell indicates that a subsequent increase in lymphocyte populations is likely.
Once the APCs have been activated, antigens are presented CD8+ and CD4+ T
cells that proliferate and are subsequently recruited back to the treated tumor, as well
270
as, circulate systemically to target and kill distal tumor cells expressing the targeted
tumor specific antigens. As discussed earlier, the reduction of T regulatory cells is
important for changing the tumor microenvironment and shifting the immune response
from “cold” to “hot”, allowing these anti-tumor CD4+ and CD8+ T cells access to
malignant cells. Histotripsy has been shown to reduce the magnitude of T regulatory
cells and increase the ratio of CD8+ to T regulatory cells in both the tumor-draining
lymph nodes and spleens of treated mice (Huang et al., 2012; Qu et al., 2020). For
example, the ratio of CD8+:CD4+ cells in the spleen was found to increase after treating
melanoma tumors with CCH (Qu et al., 2020). Additionally, after treating prostate
tumors with mHIFU, the ratio of CD8+:CD4+ cells in the spleen was also found to be
increased, and protective against subsequent tumor challenges (Huang et al., 2012).
However, in treating neuroblastoma tumors with mHIFU, the ratio of CD8+:CD4+ cells in
the spleen was found to decrease (Eranki et al., 2020). This difference in the increase
versus decrease of the CD8+:CD4+ ratio is not necessarily a sign that regulatory T cells
are predominate nor is it indication of poor prognosis. This shift in ratio could indicate
helper T cells primed to increase an immune response in response to the damage that
could lead to a pro-inflammatory response. Beyond regulatory T cells, histotripsy has
been found to increase the number of CD8+ and CD4+ helper cells in the spleen (Xing
et al., 2008; Eranki et al., 2020; Qu et al., 2020). These changes in T cell populations in
secondary lymphoid organs after histotripsy treatment implies that similar changes
occur within the tumor, and that these cells are primed against tumor specific antigens.
While all of these studies established correlative changes in T cells after
histotripsy treatment, it is important to determine the efficacy and tumor specificity of
271
programmed cells. CD8+ cells isolated from the spleens of mice that had subcutaneous
colon adenocarcinoma tumors 10 days post-treatment with mHIFU were co-cultured
with the corresponding cell line (Hu et al., 2007). The CD8+ T cells were found to be
more cytotoxicity against the cancer cells compared to the lymphocytes isolated from
mice treated with tHIFU. In this same study, ELIspot assays showed that mHIFU
generated a higher magnitude of tumor-specific CD8+ cells than tHIFU (Hu et al., 2007).
In a similar study with histotripsy treated prostate tumors, CD8+ T cells harvested from
the spleen were found to be tumor-specific and were subsequently activated when
challenged in vitro with the tumor cells (Huang et al., 2012). Finally, in a third study, the
treatment of a known antigen into a cell line before tumor engraftment allowed for the
determination of CD8+ T cells in the tumor-draining lymph nodes that were primed
against tumor-specific antigens after treatment with CCH (Qu et al., 2020). Together,
these studies support the hypothesis that histotripsy treatment of the local tumor is
effective at generating tumor specific CD8+ cytotoxic T cells that are found systemically
in distal lymph nodes and the spleen.
The efficacy of adaptive immune system activation is shown through systemic
tumor killing both through the reduction of metastasis and control of contralateral
tumors. Several studies have evaluated this phenomenon. For example, in a study
using immunocompetent New Zealand White rabbits with VX-2 (leporine papilloma)
tumors grown within a single kidney, histotripsy treatment on day 13 did not show a
change in metastasis by day 19 (Styn et al., 2012). Ultimately, the authors associated
the lack of difference to the aggression of the cancer (Styn et al., 2012). Beyond this
study, it should be noted that the majority of histotripsy studies have actually shown
272
significant improvements in metastasis (Xing et al., 2008; Qu et al., 2020). For example,
in a murine melanoma model, tumors were grown subcutaneously, treated with
histotripsy, and amputated two days after treatment (Xing et al., 2008). For most studies
conducted to date, decreases in lung metastasis are correlated to systemic cancer
control and suggestive of an abscopal response. However, as an alternative
interpretation of the data, the decrease in metastasis could simply be an effect of the
primary tumor ablation not being able to produce as many circulating tumor cells due to
its reduced size. This is a common critique of focal tumor ablation studies that report
changes in metastatic burden.
Based on the current data in the field, we believe that it is likely that both
interpretations are accurate; whereby, the decrease in metastasis is due to both the
activation of the systemic anti-tumor immune response and the debulking of the primary
tumor. Thus, it is essential that future mechanistic work in the field design experiments
designed to better establish causation. For example, in addition to the prevention of
additional tumor growth through reduced metastasis, there are multiple reports of CD8+
cytotoxic T cells in contralateral tumors (Schade et al., 2019; Qu et al., 2020). To
evaluate histotripsy in these contralateral models, mice were inoculated with bilateral
subcutaneous B16GP33 melanoma tumors and after 10 days of tumor growth one of
these tumors were treated. In the treated tumor, there was a sharp infiltration of CD8+ T
cells that localized in the ablation margin, while over the course of a week the
contralateral-untreated tumor slowly saw a perfuse increase in CD8+ T cells (Qu et al.,
2020). This study establishes that in injected, identical tumors that a systemic effect can
be achieved. As a de novo model of contralateral tumors, Eker rats have been
273
deployed. Eker rats are a model with an insertion in the rat homologue of the human
tuberous sclerosis gene (TSC2) that spontaneously develop multiple renal cell
carcinoma tumors throughout both of their kidneys (Cook and Walker, 2004). In a study
of boiling histotripsy, Eker rats had one approximately 0.5cm3 tumor treated and after
48hrs, their treated and contralateral kidneys were collected for immunohistochemistry
staining for CD8+ cells. This found that if one tumor was treated then both tumor riddled
kidneys would see an influx of CD8+ T cells, while sham treated rats did not see a
change in either kidney (Schade et al., 2019). These contralateral tumors have been
reported to have decreased growth rates after histotripsy treatment of the targeted
primary tumor, and more significantly than radiation ablation or RFA (Qu et al., 2020).
Using mice with B16GP33 tumors, after 10 days of tumor growth mice were either
treated with histotripsy, RFA, radiation therapy, or sham-control treated. This study
found that the mice that were treated with histotripsy saw a significant increase in tumor
infiltrating CD8+ T cells in treated tumors compared to RFA and radiation ablation,
neither of which saw any increase from sham-controls (Qu et al., 2020). This not only
further strengthens the idea that histotripsy can stimulate a systemic immune response,
but also supports the idea that non-thermal and non-ionizing therapies are more
effective.
While there is a reduction in metastasis in these studies, the more established
contralateral tumors only have their growth slowed, not inhibited nor reversed. This
implies that while there is a systemic immune response against the cancer, it is not
strong enough to prevent growth or be curative without combination with other
therapeutic methods or additional treatments. Further, the efficacy of mHIFU treatment
274
of generating systemic tumor-specific protection has been shown with the reduction in
tumor growth rate in challenge tumors injected 6 days after treatment (Hu et al., 2007).
Notably, the effects of mHIFU to minimize the growth rates of treated tumors were more
effective than tHIFU, even though the debulking on the primary tumor was much greater
(83% and 43% reduction in tumor volume respectively) (Hu et al., 2007). This indicates
that the non-thermal ablation modality stimulated a stronger immune response.
However, as with the pre-existing contralateral tumors, the post-treatment injection
challenge still allowed for tumor growth. Together these studies reveal that histotripsy
has the power to prime the immune system sufficiently to reduce systemic tumor burden
through control of naturally occurring metastasis, spontaneous similar-primary tumors,
and secondary tumors injected as a challenge.
7.6.6 Combination Therapy with Checkpoint-Inhibitors
Given that many cancers are immunologically resistant to checkpoint-inhibitors
due, in part, to the immunological state of the tumor microenvironment, it is critical to
explore the potential of histotripsy as a strategy to improve tumor responsiveness. The
increased release of DAMPs, shift in the inflammatory state of the tumor
microenvironment, enhanced recruitment of immune cells within the tumor, and
evidence of a systemic anti-tumor immune response all discussed above indicate that
histotripsy can have local and systemic immunomodulatory effects that may be
favorable for these classes of therapeutics. Based on these changes, it is reasonable to
hypothesize that the shift of the tumor microenvironment to a “hot” environment should
275
increase the effects of checkpoint inhibitors allowing for a potentially stronger anti-tumor
immune response. The efficacy of anti-CTLA-4, anti-PD-1, and anti-PD-L1 therapies is
correlative to the levels of expression of CTLA-4, CD80/86, PD-1, and PD-L1 in the
patients’ tumor cells, APCs, and T cells (Fig.3). Therefore, before studying the effects of
specific checkpoint inhibitors, it is critical to determine if there is supporting data to
suggest that histotripsy may have a synergistic effect (Wilky, 2019). For example, anti-
PD-L1 targeted therapies are not typically used in cases where tumors do not express
high levels of PD-L1. However, this is not always a clear decision, especially for
checkpoint inhibitors that can be induced or up-regulated following treatment. Take for
instance the expression of CD80/86 on dendritic cells, which was found to increase in
vitro when stimulated by mHIFU supernatant, indicating that the lysate released from
cells after histotripsy treatment can stimulate an upregulation of a checkpoint inhibitor
(Hu et al., 2005). In vivo, CD86 was significantly increased on dendritic cells found in
the spleen, and CD80 was increased significantly on dendritic cells in both spleen and
tumor draining lymph nodes (Huang et al., 2012). It has also been shown that there is
increase in PD-L1 in tumors 72 hours after treatment with mHIFU (Eranki et al., 2020).
As mentioned above, this could be due to the downstream effects of IFNγ production
following treatment. While this is typically detrimental to the systemic immune response,
these in vivo upregulations of checkpoint molecules after histotripsy treatment actually
indicates that there may be an increased window of opportunity to utilize checkpoint
inhibitors for synergistic therapy to potentially improve anti-tumor immunity.
With the multiple studies suggesting that histotripsy ablation can increase the
expression of both CTLA-4 and PD-1 pathway receptors, it is reasonable to hypothesize
276
that histotripsy in combination with checkpoint inhibitor therapies should have a
synergistic effect. This has been explored in a pair of recent studies. In one study,
individual CTLA-4 antibody therapy and CCH ablation decreased the rate of
contralateral tumor growth, but there was an even more significant decrease with
combined therapy in both melanoma and hepatocellular carcinoma (Qu et al., 2020).
For this study, one group of mice bearing subcutaneous B16GP33 melanoma tumors
were administered two doses of CTLA-4 monoclonal antibody prior to histotripsy and an
additional treatment afterwards. Another group of mice bearing subcutaneous Hepa1-6
hepatocellular carcinoma tumors were given three doses of CTLA-4 monoclonal
antibody prior and one post histotripsy treatment. For both of these tumor studies,
melanoma and hepatocellular carcinoma, the combination of CTLA-4 and histotripsy
yielded significantly reduced tumor volume compared to either treatment individually
(Qu et al., 2020). In a second study, three doses of both CTLA-4 and PD-L1 antibodies
in the days following mHIFU treatment of subcutaneous neuroblastoma tumors
significantly increased the survival of animals compared to other therapeutic
combinations (Eranki et al., 2020). This study went further and performed a similar
experiment, but instead of only looking at the effects of a single mHIFU treated tumor
there was a second tumor grown contralaterally. In this report, after unilateral treatment
with mHIFU and systemic CTLA-4 and PD-L1 therapy there was complete remission of
both tumors, an effect that was not seen with any other therapy combination (Eranki et
al., 2020). Together, these studies demonstrate the improved therapeutic effects of the
combined therapy over either histotripsy or checkpoint inhibitors independently.
277
7.7 Conclusion and Future Outlook
As histotripsy therapies have been developed over the past decade, the
knowledge about the immune response has started to develop a more complete picture
about the mechanisms of action from DAMP and anti-tumor mediator release, to
changes in local cellular immune populations, development of a systemic immune
response, and therapeutic synergism with the inclusion of checkpoint inhibitor therapies
(Fig. 5). In sum, these studies suggest that there is a reproducible, consistent, and
perhaps even tunable immune effect generated by histotripsy modalities that is
consistent across multiple tumor types. However, while these data are certainly exciting
and potentially paradigm shifting, the number of studies in this field are still quite limited
and most are based on murine models with inconsistent tumor ablation quality. Looking
towards the future, there needs to be an effort to compare the various histotripsy
treatments and doses, as well as other mHIFU methods, in order to better understand
the relationship between the extent of ablation in stimulating an immune response.
Likewise, as these therapies begin to be utilized in human trials, it will be crucial to
translate these findings into actionable results relevant to human patients. Additional
basic studies and preclinical animal trials are also still needed to develop missing
mechanistic insight and more translationally relevant studies are needed to ensure
these findings occur outside of the typical model organisms. Despite these limitations,
the benefits of histotripsy over other thermal ablation modalities suggest the potential of
this focal tumor ablation therapy to induce a systemic anti-tumor immune response and
278
therefore supports the hypothesis that histotripsy has a high likelihood of positively
impacting the clinical outcomes for cancer patients.
7.8 Acknowledgements
This work was supported by the Virginia Maryland College of Veterinary Medicine;
The Virginia Tech Institute for Critical Technology and Applied Sciences Center for
Engineered Health; the Focused Ultrasound Foundation; and The National Institutes of
Health. The content is solely the responsibility of the authors and does not necessarily
represent the official views of the NIH or any other funding agency. All figures were
created using Biorender.com.
279
7.9 References
Adam, R., Kitano, Y., Abdelrafee, A., Allard, M.-A., and Baba, H. (2020). Debulking
surgery for colorectal liver metastases: Foolish or chance? Surgical Oncology 33,
266-269.
Antoniades, J., Brady, L.W., and Lightfoot, D.A. (1977). Lymphangiographic
demonstration of the abscopal effect in patients with malignant lymphomas.
International Journal of Radiation Oncology* Biology* Physics 2(1-2), 141-147.
Arlauckas, S.P., Garren, S.B., Garris, C.S., Kohler, R.H., Oh, J., Pittet, M.J., et al.
(2018). Arg1 expression defines immunosuppressive subsets of tumor-
associated macrophages. Theranostics 8(21), 5842.
Armengol, C., Sarrias, M.R., and Sala, M. (2018). Hepatocellular carcinoma: present
and future. Medicina Clínica (English Edition) 150(10), 390-397.
Arnaud‐Coffin, P., Maillet, D., Gan, H.K., Stelmes, J.J., You, B., Dalle, S., et al. (2019).
A systematic review of adverse events in randomized trials assessing immune
checkpoint inhibitors. International Journal of Cancer 145(3), 639-648.
Bader, K.B., Vlaisavljevich, E., and Maxwell, A.D. (2019). For whom the bubble grows:
physical principles of bubble nucleation and dynamics in histotripsy ultrasound
therapy. Ultrasound in medicine & biology.
Bastianpillai, C., Petrides, N., Shah, T., Guillaumier, S., Ahmed, H.U., and Arya, M.
(2015). Harnessing the immunomodulatory effect of thermal and non-thermal
ablative therapies for cancer treatment. Tumor Biology 36(12), 9137-9146.
280
Befeler, A.S., and Di Bisceglie, A.M. (2002). Hepatocellular carcinoma: diagnosis and
treatment. Gastroenterology 122(6), 1609-1619.
Beitel-White, N., Martin, R.C.G., Li, Y., Brock, R.M., Allen, I.C., and Davalos, R.V.
(2019). Real-time prediction of patient immune cell modulation during irreversible
electroporation therapy. Sci Rep 9(1), 17739. doi: 10.1038/s41598-019-53974-w.
Bolat, F., Kayaselcuk, F., Nursal, T., Yagmurdur, M., Bal, N., and Demirhan, B. (2006).
Microvessel density, VEGF expression, and tumor-associated macrophages in
breast tumors: correlations with prognostic parameters. Vascular 14, 15.
Cao, X., Cai, S.F., Fehniger, T.A., Song, J., Collins, L.I., Piwnica-Worms, D.R., et al.
(2007). Granzyme B and perforin are important for regulatory T cell-mediated
suppression of tumor clearance. Immunity 27(4), 635-646.
Capps, J., and Smith, J. (1907). Experiments on the Leukolytic Action of the Blood
Serum of Cases of Leukaemia Treated with X-Ray and the Injection of Human
Leukolytic Serum in a Case of Leukaemia. The Journal of experimental medicine
9(1), 51-63.
Chandra, D., Jahangir, A., Cornelis, F., Rombauts, K., Meheus, L., Jorcyk, C.L., et al.
(2016). Cryoablation and Meriva have strong therapeutic effect on triple-negative
breast cancer. Oncoimmunology 5(1), e1049802. doi:
10.1080/2162402x.2015.1049802.
Chen, F., Bao, H., Xie, H., Tian, G., and Jiang, T. (2019). Heat shock protein expression
and autophagy after incomplete thermal ablation and their correlation.
International Journal of Hyperthermia 36(1), 95-103.
281
Chiorean, E.G., and Coveler, A.L. (2015). Pancreatic cancer: optimizing treatment
options, new, and emerging targeted therapies. Drug Des Devel Ther 9, 3529-
3545. doi: 10.2147/DDDT.S60328.
Chung, A.C., and Lan, H.Y. (2011). Chemokines in renal injury. Journal of the American
Society of Nephrology 22(5), 802-809.
Coad, J.E., Kosari, K., Humar, A., and Sielaff, T.D. (2003). Radiofrequency ablation
causes ‘thermal fixation’of hepatocellular carcinoma: a post‐liver transplant
histopathologic study. Clinical transplantation 17(4), 377-384.
Cook, J.D., and Walker, C.L. (2004). The Eker rat: establishing a genetic paradigm
linking renal cell carcinoma and uterine leiomyoma. Curr Mol Med 4(8), 813-824.
doi: 10.2174/1566524043359656.
Curiel, T.J., Coukos, G., Zou, L., Alvarez, X., Cheng, P., Mottram, P., et al. (2004).
Specific recruitment of regulatory T cells in ovarian carcinoma fosters immune
privilege and predicts reduced survival. Nature medicine 10(9), 942-949.
Dai, Z., Wang, Z., Lei, K., Liao, J., Peng, Z., Lin, M., et al. (2021). Irreversible
electroporation induces CD8+ T cell immune response against post-ablation
hepatocellular carcinoma growth. Cancer Letters. doi:
https://doi.org/10.1016/j.canlet.2021.01.001.
Dan, J., Zhang, Y., Peng, Z., Huang, J., Gao, H., Xu, L., et al. (2013). Postoperative
neutrophil-to-lymphocyte ratio change predicts survival of patients with small
hepatocellular carcinoma undergoing radiofrequency ablation. PloS one 8(3),
e58184.
282
Demaria, S., Ng, B., Devitt, M.L., Babb, J.S., Kawashima, N., Liebes, L., et al. (2004).
Ionizing radiation inhibition of distant untreated tumors (abscopal effect) is
immune mediated. Int J Radiat Oncol Biol Phys 58(3), 862-870. doi:
10.1016/j.ijrobp.2003.09.012.
den Brok, M.H., Sutmuller, R.P., van der Voort, R., Bennink, E.J., Figdor, C.G., Ruers,
T.J., et al. (2004). In situ tumor ablation creates an antigen source for the
generation of antitumor immunity. Cancer research 64(11), 4024-4029.
Domingo-Musibay, E., Heun, J.M., Nevala, W.K., Callstrom, M., Atwell, T., Galanis, E.,
et al. (2017). Endogenous Heat-Shock Protein Induction with or Without
Radiofrequency Ablation or Cryoablation in Patients with Stage IV Melanoma.
Oncologist 22(9), 1026-e1093. doi: 10.1634/theoncologist.2017-0060.
Dromi, S.A., Walsh, M.P., Herby, S., Traughber, B., Xie, J., Sharma, K.V., et al. (2009).
Radiofrequency ablation induces antigen-presenting cell infiltration and
amplification of weak tumor-induced immunity. Radiology 251(1), 58-66.
Eranki, A., Srinivasan, P., Ries, M., Kim, A., Lazarski, C.A., Rossi, C.T., et al. (2020).
High-Intensity Focused Ultrasound (HIFU) Triggers Immune Sensitization of
Refractory Murine Neuroblastoma to Checkpoint Inhibitor Therapy. Clinical
Cancer Research 26(5), 1152-1161.
Feng, L., Qi, Q., Wang, P., Chen, H., Chen, Z., Meng, Z., et al. (2018). Serum levels of
IL-6, IL-8, and IL-10 are indicators of prognosis in pancreatic cancer. Journal of
International Medical Research 46(12), 5228-5236.
283
Fleming, V., Hu, X., Weber, R., Nagibin, V., Groth, C., Altevogt, P., et al. (2018).
Targeting myeloid-derived suppressor cells to bypass tumor-induced
immunosuppression. Frontiers in immunology 9, 398.
Fodor, P., White, B., and Khan, R. (2020). Inflammation-The role of ATP in pre-
eclampsia. Microcirculation 27(1), e12585. doi: 10.1111/micc.12585.
Fridlender, Z.G., Sun, J., Kim, S., Kapoor, V., Cheng, G., Ling, L., et al. (2009).
Polarization of tumor-associated neutrophil phenotype by TGF-β:“N1” versus
“N2” TAN. Cancer cell 16(3), 183-194.
Fu, L.-Q., Du, W.-L., Cai, M.-H., Yao, J.-Y., Zhao, Y.-Y., and Mou, X.-Z. (2020). The
roles of tumor-associated macrophages in tumor angiogenesis and metastasis.
Cellular Immunology, 104119.
Gajewski, T.F. (Year). "The next hurdle in cancer immunotherapy: Overcoming the non–
T-cell–inflamed tumor microenvironment", in: Seminars in oncology: Elsevier),
663-671.
Gao, X., Sui, H., Zhao, S., Gao, X., Su, Y., and Qu, P. (2020). Immunotherapy
Targeting Myeloid-Derived Suppressor Cells (MDSCs) in Tumor
Microenvironment. Front Immunol 11, 585214. doi: 10.3389/fimmu.2020.585214.
Gong, T., Liu, L., Jiang, W., and Zhou, R. (2019). DAMP-sensing receptors in sterile
inflammation and inflammatory diseases. Nature Reviews Immunology, 1-18.
Gunz, F. (1953). Bone Marrow Changes in Patients with Chronic Leukemia Treated by
Splenic X-irradiation: Preliminary Report. Blood 8(8), 687-692.
Hajizadeh, F., Maleki, L.A., Alexander, M., Mikhailova, M.V., Masjedi, A., Ahmadpour,
M., et al. (2020). Tumor-associated neutrophils as new players in
284
immunosuppressive process of the tumor microenvironment in breast cancer.
Life sciences, 118699.
Hall, T.L., Hempel, C.R., Wojno, K., Xu, Z., Cain, C.A., and Roberts, W.W. (2009).
Histotripsy of the prostate: dose effects in a chronic canine model. Urology 74(4),
932-937.
Harris, H.E., and Andersson, U. (2004). Mini‐review: the nuclear protein HMGB1 as a
proinflammatory mediator. European journal of immunology 34(6), 1503-1512.
Hempel, C.R., Hall, T.L., Cain, C.A., Fowlkes, J.B., Xu, Z., and Roberts, W.W. (2011).
Histotripsy fractionation of prostate tissue: local effects and systemic response in
a canine model. The Journal of urology 185(4), 1484-1489.
Henry, C.J., Ornelles, D.A., Mitchell, L.M., Brzoza-Lewis, K.L., and Hiltbold, E.M.
(2008). IL-12 produced by dendritic cells augments CD8+ T cell activation
through the production of the chemokines CCL1 and CCL17. J Immunol 181(12),
8576-8584. doi: 10.4049/jimmunol.181.12.8576.
Hotchkiss, D.J., and Block, M.H. (1962). Effect of splenic irradiation on systemic
hematopoiesis. Archives of internal medicine 109(6), 695-711.
Hsieh, J.J., Purdue, M.P., Signoretti, S., Swanton, C., Albiges, L., Schmidinger, M., et
al. (2017). Renal cell carcinoma. Nature reviews Disease primers 3, 17009.
Hu, J.M., Liu, K., Liu, J.H., Jiang, X.L., Wang, X.L., Yang, L., et al. (2017). The
increased number of tumor-associated macrophage is associated with
overexpression of VEGF-C, plays an important role in Kazakh ESCC invasion
and metastasis. Experimental and molecular pathology 102(1), 15-21.
285
Hu, Z., Yang, X.Y., Liu, Y., Morse, M.A., Lyerly, H.K., Clay, T.M., et al. (2005). Release
of endogenous danger signals from HIFU-treated tumor cells and their
stimulatory effects on APCs. Biochemical and biophysical research
communications 335(1), 124-131.
Hu, Z., Yang, X.Y., Liu, Y., Sankin, G.N., Pua, E.C., Morse, M.A., et al. (2007).
Investigation of HIFU-induced anti-tumor immunity in a murine tumor model.
Journal of translational medicine 5(1), 34.
Huang, X., Yuan, F., Liang, M., Lo, H.-W., Shinohara, M.L., Robertson, C., et al. (2012).
M-HIFU inhibits tumor growth, suppresses STAT3 activity and enhances tumor
specific immunity in a transplant tumor model of prostate cancer. PloS one 7(7),
e41632.
Ito, F., Vardam, T.D., Appenheimer, M.M., Eng, K.H., Gollnick, S.O., Muhitch, J.B., et al.
(2019). In situ thermal ablation augments antitumor efficacy of adoptive T cell
therapy. International Journal of Hyperthermia 36(sup1), 22-36.
Jiang, H., Hegde, S., and DeNardo, D.G. (2017). Tumor-associated fibrosis as a
regulator of tumor immunity and response to immunotherapy. Cancer
Immunology, Immunotherapy 66(8), 1037-1048.
Kalathil, S., Lugade, A.A., Miller, A., Iyer, R., and Thanavala, Y. (2013). Higher
frequencies of GARP+ CTLA-4+ Foxp3+ T regulatory cells and myeloid-derived
suppressor cells in hepatocellular carcinoma patients are associated with
impaired T-cell functionality. Cancer research 73(8), 2435-2444.
286
Kaminski, J.M., Shinohara, E., Summers, J.B., Niermann, K.J., Morimoto, A., and
Brousal, J. (2005). The controversial abscopal effect. Cancer treatment reviews
31(3), 159-172.
Keisari, Y. (2017). Tumor abolition and antitumor immunostimulation by physico-
chemical tumor ablation. Front Biosci (Landmark Ed) 22, 310-347.
Kepp, O., Marabelle, A., Zitvogel, L., and Kroemer, G. (2019). Oncolysis without
viruses—inducing systemic anticancer immune responses with local therapies.
Nature Reviews Clinical Oncology, 1-16.
Khokhlova, V.A., Fowlkes, J.B., Roberts, W.W., Schade, G.R., Xu, Z., Khokhlova, T.D.,
et al. (2015). Histotripsy methods in mechanical disintegration of tissue: Towards
clinical applications. International journal of hyperthermia 31(2), 145-162.
Kim, H., Park, B.K., and Kim, C.K. (2008). Spontaneous regression of pulmonary and
adrenal metastases following percutaneous radiofrequency ablation of a
recurrent renal cell carcinoma. Korean journal of radiology 9(5), 470.
Kindlund, B., Sjöling, Å., Yakkala, C., Adamsson, J., Janzon, A., Hansson, L.E., et al.
(2017). CD4(+) regulatory T cells in gastric cancer mucosa are proliferating and
express high levels of IL-10 but little TGF-β. Gastric Cancer 20(1), 116-125. doi:
10.1007/s10120-015-0591-z.
Ko, J.S., Zea, A.H., Rini, B.I., Ireland, J.L., Elson, P., Cohen, P., et al. (2009). Sunitinib
mediates reversal of myeloid-derived suppressor cell accumulation in renal cell
carcinoma patients. Clin Cancer Res 15(6), 2148-2157. doi: 10.1158/1078-
0432.Ccr-08-1332.
287
Kondo, S., Okusaka, T., Ueno, H., Ikeda, M., and Morizane, C. (2006). Spontaneous
regression of hepatocellular carcinoma. International journal of clinical oncology
11(5), 407-411.
Korbelik, M., Banáth, J., Saw, K.M., Zhang, W., and Čiplys, E. (2015). Calreticulin as
cancer treatment adjuvant: combination with photodynamic therapy and
photodynamic therapy-generated vaccines. Frontiers in oncology 5, 15.
Kratochvill, F., Neale, G., Haverkamp, J.M., Van de Velde, L.-A., Smith, A.M.,
Kawauchi, D., et al. (2015). TNF counterbalances the emergence of M2 tumor
macrophages. Cell reports 12(11), 1902-1914.
Ladoire, S., Martin, F., and Ghiringhelli, F. (2011). Prognostic role of FOXP3+ regulatory
T cells infiltrating human carcinomas: the paradox of colorectal cancer. Cancer
Immunology, Immunotherapy 60(7), 909-918.
Lake, A., Xu, Z., Wilkinson, J., Cain, C., and Roberts, W. (2008). Renal ablation by
histotripsy—does it spare the collecting system? The Journal of urology 179(3),
1150-1154.
Lang, R., Patel, D., Morris, J.J., Rutschman, R.L., and Murray, P.J. (2002). Shaping
gene expression in activated and resting primary macrophages by IL-10. The
Journal of Immunology 169(5), 2253-2263.
Lapeyre-Prost, A., Terme, M., Pernot, S., Pointet, A.-L., Voron, T., Tartour, E., et al.
(2017). "Immunomodulatory activity of VEGF in cancer," in International review of
cell and molecular biology. Elsevier), 295-342.
288
Leuchte, K., Staib, E., Thelen, M., Gödel, P., Lechner, A., Zentis, P., et al. (2020).
Microwave ablation enhances tumor-specific immune response in patients with
hepatocellular carcinoma. Cancer Immunology, Immunotherapy, 1-15.
Levings, M.K., Bacchetta, R., Schulz, U., and Roncarolo, M.G. (2002). The role of IL-10
and TGF-β in the differentiation and effector function of T regulatory cells.
International archives of allergy and immunology 129(4), 263-276.
Lin, K.W., Kim, Y., Maxwell, A.D., Wang, T.Y., Hall, T.L., Xu, Z., et al. (2014). Histotripsy
beyond the intrinsic cavitation threshold using very short ultrasound pulses:
microtripsy. IEEE Trans Ultrason Ferroelectr Freq Control 61(2), 251-265. doi:
10.1109/TUFFC.2014.6722611.
Liu, S., Zhang, C., Wang, B., Zhang, H., Qin, G., Li, C., et al. (2021). Regulatory T cells
promote glioma cell stemness through TGF-β-NF-κB-IL6-STAT3 signaling.
Cancer Immunol Immunother. doi: 10.1007/s00262-021-02872-0.
Lu, P., Zhu, X.-Q., Xu, Z.-L., Zhou, Q., Zhang, J., and Wu, F. (2009). Increased
infiltration of activated tumor-infiltrating lymphocytes after high intensity focused
ultrasound ablation of human breast cancer. Surgery 145(3), 286-293.
Mabuchi, S., Sasano, T., and Komura, N. (2021). Targeting Myeloid-Derived
Suppressor Cells in Ovarian Cancer. Cells 10(2). doi: 10.3390/cells10020329.
Mimura, K., Teh, J.L., Okayama, H., Shiraishi, K., Kua, L.F., Koh, V., et al. (2018). PD-
L1 expression is mainly regulated by interferon gamma associated with JAK-
STAT pathway in gastric cancer. Cancer Sci 109(1), 43-53. doi:
10.1111/cas.13424.
289
Minami, Y., Nishida, N., and Kudo, M. (2019). Radiofrequency ablation of liver
metastasis: potential impact on immune checkpoint inhibitor therapy. European
radiology 29(9), 5045-5051.
Najafi, M., Hashemi Goradel, N., Farhood, B., Salehi, E., Nashtaei, M.S., Khanlarkhani,
N., et al. (2019). Macrophage polarity in cancer: A review. Journal of cellular
biochemistry 120(3), 2756-2765.
Ni, L., and Lu, J. (2018). Interferon gamma in cancer immunotherapy. Cancer medicine
7(9), 4509-4516.
Nobler, M.P. (1969). The abscopal effect in malignant lymphoma and its relationship to
lymphocyte circulation. Radiology 93(2), 410-412.
Ostuni, R., Kratochvill, F., Murray, P.J., and Natoli, G. (2015). Macrophages and cancer:
from mechanisms to therapeutic implications. Trends in immunology 36(4), 229-
239.
Pahk, K.J., Shin, C.-H., Bae, I.Y., Yang, Y., Kim, S.-H., Pahk, K., et al. (2019). Boiling
Histotripsy-induced Partial Mechanical Ablation Modulates Tumour
Microenvironment by Promoting Immunogenic Cell Death of Cancers. Scientific
reports 9(1), 1-12.
Pandit, H., Hong, Y.K., Li, Y., Rostas, J., Pulliam, Z., Li, S.P., et al. (2019). Evaluating
the regulatory immunomodulation effect of irreversible electroporation (IRE) in
pancreatic adenocarcinoma. Annals of surgical oncology 26(3), 800-806.
Pang, L., Han, S., Jiao, Y., Jiang, S., He, X., and Li, P. (2017). Bu Fei Decoction
attenuates the tumor associated macrophage stimulated proliferation, migration,
290
invasion and immunosuppression of non-small cell lung cancer, partially via IL-10
and PD-L1 regulation. International Journal of Oncology 51(1), 25-38.
Pillai, K., Akhter, J., Chua, T.C., Shehata, M., Alzahrani, N., Al-Alem, I., et al. (2015).
Heat sink effect on tumor ablation characteristics as observed in monopolar
radiofrequency, bipolar radiofrequency, and microwave, using ex vivo calf liver
model. Medicine 94(9).
Pisetsky, D.S. (2011). Cell death in the pathogenesis of immune-mediated diseases: the
role of HMGB1 and DAMP-PAMP complexes. Swiss medical weekly 141,
w13256.
Plate, J. (2011). Clinical trials of vaccines for immunotherapy in pancreatic cancer.
Expert review of vaccines 10(6), 825-836.
Qu, S., Worlikar, T., Felsted, A.E., Ganguly, A., Beems, M.V., Hubbard, R., et al. (2020).
Non-thermal histotripsy tumor ablation promotes abscopal immune responses
that enhance cancer immunotherapy. Journal for ImmunoTherapy of Cancer
8(1).
Ringel-Scaia, V.M., Beitel-White, N., Lorenzo, M.F., Brock, R.M., Huie, K.E.,
Coutermarsh-Ott, S., et al. (2019). High-frequency irreversible electroporation is
an effective tumor ablation strategy that induces immunologic cell death and
promotes systemic anti-tumor immunity. EBioMedicine 44, 112-125. doi:
10.1016/j.ebiom.2019.05.036.
Rodriguez, P.C., Ernstoff, M.S., Hernandez, C., Atkins, M., Zabaleta, J., Sierra, R., et al.
(2009). Arginase I-producing myeloid-derived suppressor cells in renal cell
291
carcinoma are a subpopulation of activated granulocytes. Cancer Res 69(4),
1553-1560. doi: 10.1158/0008-5472.Can-08-1921.
Saccomandi, P., Lapergola, A., Longo, F., Schena, E., and Quero, G. (2018). Thermal
ablation of pancreatic cancer: A systematic literature review of clinical practice
and pre-clinical studies. International Journal of Hyperthermia 35(1), 398-418.
Salama, P., Phillips, M., Grieu, F., Morris, M., Zeps, N., Joseph, D., et al. (2009).
Tumor-infiltrating FOXP3+ T regulatory cells show strong prognostic significance
in colorectal cancer. Journal of clinical oncology 27(2), 186-192.
Sanchez-Ortiz, R.F., Tannir, N., Ahrar, K., and Wood, C.G. (2003). Spontaneous
regression of pulmonary metastases from renal cell carcinoma after radio
frequency ablation of primary tumor: in situ tumor vaccine? The Journal of
urology 170(1), 178-179.
Schade, G.R. (2018). Evaluation of the Systemic Response to Boiling Histotripsy
Treatment for Renal Carcinoma.
Schade, G.R., Wang, Y.-N., D'Andrea, S., Hwang, J.H., Liles, W.C., and Khokhlova,
T.D. (2019). Boiling Histotripsy Ablation of Renal Cell Carcinoma in the Eker Rat
Promotes a Systemic Inflammatory Response. Ultrasound in medicine & biology
45(1), 137-147.
Scheffer, H.J., Stam, A.G., Geboers, B., Vroomen, L.G., Ruarus, A., de Bruijn, B., et al.
(2019). Irreversible electroporation of locally advanced pancreatic cancer
transiently alleviates immune suppression and creates a window for antitumor T
cell activation. Oncoimmunology 8(11), 1652532.
292
Senn, N. (1903). Case of Splenomedullary Leukaemia Successfully Treated by the Use
of the Roentgen. Med Rec 64, 281-282.
Shadhu, K., and Xi, C. (2019). Inflammation and pancreatic cancer: An updated review.
Saudi J Gastroenterol 25(1), 3-13. doi: 10.4103/sjg.SJG_390_18.
Shao, Q., O'Flanagan, S., Lam, T., Roy, P., Pelaez, F., Burbach, B.J., et al. (2019).
Engineering T cell response to cancer antigens by choice of focal therapeutic
conditions. International Journal of Hyperthermia 36(1), 130-138.
Smolock AR, C.M., Vlaisavljevich E, Gendron-Fitzpatrick A, Green C, Ziemlewicz TJ
and Lee FT (2018). Robotically Assisted Sonic Therapy as a Non-Invasive Non-
Thermal Ablation Modality: Proof of Concept in a Porcine Liver Model. Radiology
287(2), 485-493.
Smolock, A.R., Cristescu, M.M., Vlaisavljevich, E., Gendron-Fitzpatrick, A., Green, C.,
Cannata, J., et al. (2018). Robotically Assisted Sonic Therapy as a Noninvasive
Nonthermal Ablation Modality: Proof of Concept in a Porcine Liver Model.
Radiology 287(2), 485-493. doi: 10.1148/radiol.2018171544.
Styn, N.R., Hall, T.L., Fowlkes, J.B., Cain, C.A., and Roberts, W.W. (2012). Histotripsy
of renal implanted VX-2 tumor in a rabbit model: investigation of metastases.
Urology 80(3), 724-729.
Summers, C., Rankin, S.M., Condliffe, A.M., Singh, N., Peters, A.M., and Chilvers, E.R.
(2010). Neutrophil kinetics in health and disease. Trends in immunology 31(8),
318-324.
Tanaka, A., and Sakaguchi, S. (2019). Targeting Treg cells in cancer immunotherapy.
European journal of immunology 49(8), 1140-1146.
293
Tesi, R. (2019). MDSC; the most important cell you have never heard of. Trends in
pharmacological sciences 40(1), 4-7.
Thanos, L., Mylona, S., Galani, P., Pomoni, M., Pomoni, A., and Koskinas, I. (2008).
Overcoming the heat-sink phenomenon: successful radiofrequency thermal
ablation of liver tumors in contact with blood vessels. Diagnostic and
Interventional Radiology 14(1), 51.
Trellakis, S., Farjah, H., Bruderek, K., Dumitru, C., Hoffmann, T., Lang, S., et al. (2011).
Peripheral blood neutrophil granulocytes from patients with head and neck
squamous cell carcinoma functionally differ from their counterparts in healthy
donors. International journal of immunopathology and pharmacology 24(3), 683-
693.
van Beijnum, J.R., Buurman, W.A., and Griffioen, A.W. (2008). Convergence and
amplification of toll-like receptor (TLR) and receptor for advanced glycation end
products (RAGE) signaling pathways via high mobility group B1 (HMGB1).
Angiogenesis 11(1), 91-99. doi: 10.1007/s10456-008-9093-5.
van Raam, B.J., Drewniak, A., Groenewold, V., van den Berg, T.K., and Kuijpers, T.W.
(2008). Granulocyte colony-stimulating factor delays neutrophil apoptosis by
inhibition of calpains upstream of caspase-3. Blood, The Journal of the American
Society of Hematology 112(5), 2046-2054.
Vizio, B., Novarino, A., Giacobino, A., Cristiano, C., Prati, A., Ciuffreda, L., et al. (2012).
Potential plasticity of T regulatory cells in pancreatic carcinoma in relation to
disease progression and outcome. Experimental and therapeutic medicine 4(1),
70-78.
294
Vlaisavljevich, E., Greve, J., Cheng, X., Ives, K., Shi, J., Jin, L., et al. (2016a). Non-
invasive ultrasound liver ablation using histotripsy: Chronic study in an in vivo
rodent model. Ultrasound in medicine & biology 42(8), 1890-1902.
Vlaisavljevich, E., Kim, Y., Allen, S., Owens, G., Pelletier, S., Cain, C., et al. (2013a).
Image-guided non-invasive ultrasound liver ablation using histotripsy: Feasibility
study in an in vivo porcine model. Ultrasound in medicine & biology 39(8), 1398-
1409.
Vlaisavljevich, E., Kim, Y., Owens, G., Roberts, W., Cain, C., and Xu, Z. (2013b).
Effects of tissue mechanical properties on susceptibility to histotripsy-induced
tissue damage. Physics in Medicine & Biology 59(2), 253.
Vlaisavljevich, E., Kim, Y., Owens, G., Roberts, W., Cain, C., and Xu, Z. (2014a).
Effects of tissue mechanical properties on susceptibility to histotripsy-induced
tissue damage. Physics in Medicine and Biology 59(2), 253-270. doi:
10.1088/0031-9155/59/2/253.
Vlaisavljevich, E., Lin, K.W., Maxwell, A., Warnez, M., Mancia, L., Singh, R., et al.
(2015a). Effects of Ultrasound Frequency and Tissue Stiffness on the Histotripsy
Intrinsic Threshold for Cavitation. Ultrasound Med Biol.
Vlaisavljevich, E., Maxwell, A., Mancia, L., Johnsen, E., Cain, C., and Xu, Z. (2016b).
Visualizing the Histotripsy Process: Bubble Cloud-Cancer Cell Interactions in a
Tissue-Mimicking Environment. Ultrasound Med Biol. doi: S0301-5629(16)30099-
0 [pii] 10.1016/j.ultrasmedbio.2016.05.018.
Vlaisavljevich, E., Maxwell, A., Warnez, M., Johnsen, E., Cain, C., and Xu, Z. (2014b).
Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
295
mechanical properties. IEEE Transactions on Ultrasonics, Ferroelectrics, and
Frequency Control 61(2), 341-352. doi: 10.1109/TUFFC.2014.6722618.
Vlaisavljevich, E., Owens, G., Lundt, J., Teofilovic, D., Ives, K., Duryea, A., et al. (2017).
Non-invasive liver ablation using histotripsy: preclinical safety study in an in vivo
porcine model. Ultrasound in medicine & biology 43(6), 1237-1251.
Vlaisavljevich, E., Xu, Z., Arvidson, A., Jin, L., Roberts, W., and Cain, C. (2015b).
Effects of thermal preconditioning on tissue susceptibility to histotripsy.
Ultrasound in medicine & biology 41(11), 2938-2954.
Wan, S., Zhao, E., Kryczek, I., Vatan, L., Sadovskaya, A., Ludema, G., et al. (2014).
Tumor-associated macrophages produce interleukin 6 and signal via STAT3 to
promote expansion of human hepatocellular carcinoma stem cells.
Gastroenterology 147(6), 1393-1404.
Wang, D., An, G., Xie, S., Yao, Y., and Feng, G. (2016). The clinical and prognostic
significance of CD14+ HLA-DR−/low myeloid-derived suppressor cells in
hepatocellular carcinoma patients receiving radiotherapy. Tumor Biology 37(8),
10427-10433.
Wang, D., Zhang, X., Gao, Y., Cui, X., Yang, Y., Mao, W., et al. (2020). Research
Progress and Existing Problems for Abscopal Effect. Cancer Manag Res 12,
6695-6706. doi: 10.2147/cmar.S245426.
Wang, X., and Lin, Y. (2008). Tumor necrosis factor and cancer, buddies or foes? 1.
Acta Pharmacologica Sinica 29(11), 1275-1288.
296
Watford, W.T., Moriguchi, M., Morinobu, A., and O'Shea, J.J. (2003). The biology of IL-
12: coordinating innate and adaptive immune responses. Cytokine Growth Factor
Rev 14(5), 361-368. doi: 10.1016/s1359-6101(03)00043-1.
Widenmeyer, M., Shebzukhov, Y., Haen, S.P., Schmidt, D., Clasen, S., Boss, A., et al.
(2011). Analysis of tumor antigen‐specific T cells and antibodies in cancer
patients treated with radiofrequency ablation. International Journal of Cancer
128(11), 2653-2662.
Wilky, B.A. (2019). Immune checkpoint inhibitors: the linchpins of modern
immunotherapy. Immunological reviews 290(1), 6-23.
Wislez, M., Fleury-Feith, J., Rabbe, N., Moreau, J., Cesari, D., Milleron, B., et al. (2001).
Tumor-derived granulocyte-macrophage colony-stimulating factor and
granulocyte colony-stimulating factor prolong the survival of neutrophils
infiltrating bronchoalveolar subtype pulmonary adenocarcinoma. The American
journal of pathology 159(4), 1423-1433.
Wissniowski, T.T., Hänsler, J., Neureiter, D., Frieser, M., Schaber, S., Esslinger, B., et
al. (2003). Activation of tumor-specific T lymphocytes by radio-frequency ablation
of the VX2 hepatoma in rabbits. Cancer research 63(19), 6496-6500.
Worlikar, T., Mendiratta-Lala, M., Greve, J., Cho, C., Hubbard, R., Vlaisavljevich, E., et
al. (2019). "Non-Invasive Orthotopic Liver Tumor Ablation Using Histotripsy in an
in Vivo Rodent Hepatocellular Carcinoma (HCC) Model", in: Society of
Interventional Oncology. (Boston, MA, USA).
Worlikar, T., Vlaisavljevich, E., Gerhardson, T., Greve, J., Wan, S., Kuruvilla, S., et al.
(2018). Histotripsy for Non-Invasive Ablation of Hepatocellular Carcinoma (HCC)
297
Tumor in a Subcutaneous Xenograft Murine Model. Conf Proc IEEE Eng Med
Biol Soc 2018, 6064-6067. doi: 10.1109/EMBC.2018.8513650.
Wu, F., Wang, Z.-B., Lu, P., Xu, Z.-L., Chen, W.-Z., Zhu, H., et al. (2004). Activated anti-
tumor immunity in cancer patients after high intensity focused ultrasound
ablation. Ultrasound in medicine & biology 30(9), 1217-1222.
Wu, K., Lin, K., Li, X., Yuan, X., Xu, P., Ni, P., et al. (2020). Redefining Tumor-
Associated Macrophage Subpopulations and Functions in the Tumor
Microenvironment. Front Immunol 11, 1731. doi: 10.3389/fimmu.2020.01731.
Wu, L., Saxena, S., Awaji, M., and Singh, R.K. (2019). Tumor-associated neutrophils in
cancer: going pro. Cancers 11(4), 564.
Wu, L., and Zhang, X.H. (2020). Tumor-Associated Neutrophils and Macrophages-
Heterogenous but Not Chaotic. Front Immunol 11, 553967. doi:
10.3389/fimmu.2020.553967.
Xie, C., Duffy, A.G., Mabry‐Hrones, D., Wood, B., Levy, E., Krishnasamy, V., et al.
(2019). Tremelimumab in combination with microwave ablation in patients with
refractory biliary tract cancer. Hepatology 69(5), 2048-2060.
Xing, Y., Lu, X., Pua, E.C., and Zhong, P. (2008). The effect of high intensity focused
ultrasound treatment on metastases in a murine melanoma model. Biochemical
and biophysical research communications 375(4), 645-650.
Yang, R., Reilly, C.R., Rescorla, F.J., Sanghvi, N.T., Fry, F.J., Franklin Jr, T.D., et al.
(1992). Effects of high-intensity focused ultrasound in the treatment of
experimental neuroblastoma. Journal of pediatric surgery 27(2), 246-251.
298
Yin, J., Dong, J., Gao, W., and Wang, Y. (2018). A case report of remarkable response
to association of radiofrequency ablation with subsequent Atezolizumab in stage
IV nonsmall cell lung cancer. Medicine 97(44).
Youn, J.I., and Gabrilovich, D.I. (2010). The biology of myeloid‐derived suppressor cells:
the blessing and the curse of morphological and functional heterogeneity.
European journal of immunology 40(11), 2969-2975.
Zamarron, B.F., and Chen, W. (2011). Dual roles of immune cells and their factors in
cancer development and progression. International journal of biological sciences
7(5), 651.
Zhou, H., Jiang, M., Yuan, H., Ni, W., and Tai, G. (2021). Dual roles of myeloid-derived
suppressor cells induced by Toll-like receptor signaling in cancer. Oncol Lett
21(2), 149. doi: 10.3892/ol.2020.12410.
Zhou, K., Cheng, T., Zhan, J., Peng, X., Zhang, Y., Wen, J., et al. (2020). Targeting
tumor-associated macrophages in the tumor microenvironment. Oncol Lett 20(5),
234. doi: 10.3892/ol.2020.12097.
299
Chapter 8. Histotripsy Ablation Alters the Tumor Microenvironment and Promotes Immune System Activation in a Subcutaneous Model of Pancreatic
Cancer
Alissa Hendricks-Wenger, Jaqueline Sereno, Jessica Gannon, Allison Zeher, Rebecca M Brock, Natalie Betiel-White, Alexander Simon, Rafael V. Davalos, Sheryl
Coutermarsh-Ott, Eli Vlaisavljevich, Irving C. Allen
This chapter is excerpted from a manuscript published in IEEE Transactions in UFFC.
2021 IEEE. Reprinted, with permission, from: A. Hendricks-Wenger et al., "Histotripsy Ablation Alters the Tumor Microenvironment and Promotes Immune System Activation in a Subcutaneous Model of Pancreatic Cancer," in IEEE Transactions on Ultrasonics,
Ferroelectrics, and Frequency Control.
300
8.1 Abstract
Pancreatic cancer is a significant cause of cancer-related deaths in the United States
with an abysmal 5-year overall survival rate that is under 9%. Reasons for this mortality
include the lack of late-stage treatment options and the immunosuppressive tumor
microenvironment. Histotripsy is an ultrasound-guided, non-invasive, non-thermal tumor
ablation therapy that mechanically lyses targeted cells. To study the effects of
histotripsy on pancreatic cancer, we utilized an in vitro model of pancreatic
adenocarcinoma and compared the release of potential antigens following histotripsy
treatment to other ablation modalities. Histotripsy was found to release immune-
stimulating molecules at magnitudes similar to other non-thermal ablation modalities
and superior to thermal ablation modalities, which corresponded to increased innate
immune system activation in vivo. In subsequent in vivo studies, murine Pan02 tumors
were grown in mice and treated with histotripsy. Flow cytometry and rtPCR were used
to determine changes in the tumor microenvironment over time compared to untreated
animals. In mice with pancreatic tumors, we observed significantly increased tumor-
progression-free and general survival, with increased activation of the innate immune
system 24 hours post-treatment and decreased tumor-associated immune cell
populations within 14 days of treatment. This study demonstrates the feasibility of using
histotripsy for pancreatic cancer ablation and provides mechanistic insight into the initial
innate immune system activation following treatment. Further work is needed to
301
establish the mechanisms behind the immunomodulation of the tumor
microenvironment and immune effects.
8.2 Keywords
Biological Effects & Dosimetry, Therapeutics
302
8.3 Introduction
Pancreatic cancer is the fourth leading cause of cancer-related deaths, with a 9%
survival rate due to its late diagnosis and lack of curative treatment options [1, 2].
Standard treatment options are limited and include surgery, chemotherapy, and
radiation [3]. Only 20% of patients have tumors that can be surgically removed, and of
those patients the cure rate is less than 25% [4]. Ablation procedures can act as a
replacement for, or as an adjuvant to, surgery. The most commonly used ablation
modalities for treating cancers in abdominal organs are radiofrequency ablation (RFA),
microwave ablation, high-intensity focused ultrasound, and cryoablation. RFA is a
thermal, minimally-invasive ablation modality that utilizes high-frequency alternating
currents to thermally induce thermal necrosis [5]. Microwave ablation, also thermal and
minimally-invasive, induces cell death through electromagnetic microwaves that
produce friction and heat [6]. High-intensity focused ultrasound ablates cells by
depositing ultrasound energy at a focal point to rapidly increase tissue temperature [7].
Cryoablation results in tumor cell destruction through ice-crystal formation as a result of
liquid nitrogen or argon gas delivered to the tissue and does not result in protein
denaturation, which has led to increased reports of immunological effects compared to
RFA and microwave ablation [5]. The thermal ablation modalities, although efficacious
in treating certain malignancies, have not yet been ubiquitously accepted into clinics for
pancreatic cancer therapy due to the risk of thermal damage to healthy pancreatic
tissue, vasculature, and other critical structures.
303
Recent advancements have established non-thermal therapies that have the
potential to treat pancreatic cancer. For example, irreversible electroporation (IRE)
utilizes short, high-voltage electrical pulses that non-thermally open micro-pores in cell
plasma membranes, inducing cell death [8]. Clinical trials have shown that IRE can
ablate pancreatic tumors without damaging nearby critical structures and have had
dramatic effects on patient survival that may be due to the induction of
immunomodulatory mechanisms [9-14]. Given IRE’s controlled cell death mechanisms,
the procedure has been found to release immunostimulatory molecules and lead to a
more immunologically active tumor microenvironment after treatment [15]. Furthermore,
recent studies comparing the thermal ablation modalities, cryoablation, and IRE have
found that the non-thermal modalities are more potent at stimulating an anti-tumor
microenvironment [16]. These procedures still involve surgical incisions that increase
the possibility of surgery-related injury or infection.
To address the clinical limitations of IRE and other ablation modalities, new
focused ultrasound ablation methods have been studied as a completely non-invasive,
non-thermal alternative. Histotripsy is a non-thermal, non-ionizing, image-guided
ablation modality that uses focused-ultrasound to initiate acoustic cavitation, which
leads to the lysis of cells contained in the targeted area [17, 18]. Ablation of internal
targets with histotripsy is effective with little to no off-target effects [19-21]. Early studies
with histotripsy ablation of melanoma, hepatocellular carcinoma, renal cell carcinomas,
colorectal carcinomas, and neuroblastomas established that there is an activation of
local, cellular, and systemic immune responses [22-27].
304
In this study, we assess the ability of histotripsy to ablate subcutaneous
pancreatic tumors and determine the immunological changes within the treated tumors
over time. First, we compared the release of damage-associated molecular patterns
(DAMPs) known to activate the innate immune system and neoantigen generation
following histotripsy against other tumor ablation modalities. We extended these in vitro
findings using the in vivo Pan02 murine pancreatic cancer model. This mouse model
was chosen given its well characterized progression and described immune effects to
cancer therapies [15, 28, 29]. Using this model, we demonstrate effective tumor
ablation, pro-inflammatory changes in the tumor micro-environment, and identify critical
immune signaling mechanisms associated with histotripsy treatment.
8.4 Procedures
8.4.1 Tumor Injections and Monitoring
All in vivo experiments were conducted under institutional IACUC approval and
following the NIH Guide for the Care and Use of Laboratory Animals. For these studies,
male and female mice were equally utilized in the 7 to 10-week age range. Once the
entire cohort of C57/Bl6 mice reached a minimum weight of 20g, 100 µL of Pan02 cells
(DTP, DCTD Tumor Repository) at a concentration of 6.0x107 cells/mL of Matrigel
(Corning) were injected into the right flank of the mice that were anesthetized with
vaporized isoflurane (1.5 L/min oxygen flow with 1-3% isoflurane). Control animals were
injected with the same amount of Matrigel that did not contain Pan02 cells. The mice
305
were then monitored three times a week until the end of the study. Tumor diameters
were measured with calipers and calculated as the square root of two perpendicular
measurements, as previously described [24]. The weights and tumor sizes were
recorded along with the general health of the mice.
8.4.2 Histotripsy Set Up
In vivo studies used a custom 1MHz, 8-element small animal histotripsy transducer
with a geometric focus of 36 mm, an aperture size of 52.7 mm, and an f-number of 0.68.
The full-width half-maximum (FWHM) dimensions at a geometric focus of this
transducer were 0.98 mm, 0.93 mm, and 3.9 mm in transverse, elevational, and axial,
respectively. The transducer was driven via a custom high-voltage pulser designed to
generate short therapy pulses of <2 cycles controlled by a field-programmable gate
array (FPGA) board (Altera DE0-Nano Terasic Technology, Dover, DE, USA)
programmed for histotripsy therapy pulsing. The transducer was positioned in a tank of
degassed water heated to 37+4ºC beneath a custom-designed mouse surgical stage
(Fig.1A) and attached to a computer-guided 3-D positioning system with a 0.05 mm
motor resolution to control the automated volumetric treatments. A linear ultrasound
imaging probe with a frequency range of 10-18 MHz (L18-10L30H-4, Telemed,
Lithuania, EU) was coaxially aligned inside the transducer for treatment guidance and
monitoring [20, 24]. The transducer was powered by a high voltage DC power supply
(GENH750W, TDK-Lambda), and the system was controlled using a custom user-
interface operated through MATLAB (MathWorks).
306
Figure 8-1 In Vivo Experimental Setup. Murine histotripsy experiments were conducted using a 1Mhz transducer (A). Timing for treatment, euthanasia, and data collection were set as diagramed (B). Days with histology collection are noted by cassettes, flash frozen tumors noted by tubes with tumors, serum noted by tubes with serum, and flow cytometry is noted by flow charts. Tumors located with ultrasound imaging were used for planning automated ablation disks (C) and raster scan plots (D). Therapy was guided by co-axially aligned ultrasound imaging (E). Red arrow indicates a bubble cloud that was generated during treatment.
307
8.4.3 n Vitro Ablations Treatment Parameters
Pan02 cells transfected with a plasmid that produces the influenza antigen
hemagglutinin (Pan02-HA) were used for these in vitro experiments. HA is a common
surrogate used in tumor-specific antigen studies. Pan02-HA cells were ablated in four
treatments and two levels of ablation (Fig.2A). Cells were collected, centrifuged to
pellet, and were resuspended in PBS at a concentration of 10x106 cells/mL. 1 mL of
cell suspension was used for each treatment. All samples were kept on ice until
treatment. For all treatments, the partial ablation dosages were determined to be a
successful ablation with >30% viability and the full ablation dosages had <10% viability
for all samples. For each ablation modalities the prefix “f-” is added before the modality
name within the text to refer to the full ablation, and “p-” is added to refer to the partial
ablation. For example, f-histotripsy is full ablation histotripsy and p-Histotripsy is partial
ablation histotripsy.
Histotripsy treatments were done in a custom holder (Fig.2B), utilizing the same
1MHz transducer from the in vivo work at a PRF of 250Hz at a single spot for 0.5 or 5
minutes. The movement of the transducer during treatment was not necessary given the
circulation of the cell suspension caused by histotripsy that was observed during
treatment and in previous studies [30]. For cryoablation, cells were placed in liquid
nitrogen (approximately -160oC) for 30 minutes. No lower dose was used given that any
drop to therapeutically low temperatures (-20oC to -190oC) resulted in high levels of cell
death [31]. Thermal ablation consisted of cells being kept at 80oC on a heating block for
1 or 30 minutes. The protocols for cryoablation and thermal ablation were based upon a
308
previous study.[16] For samples not included in analysis, the temperatures of the
thermally treated samples were confirmed with a thermometer to reach 45oC, minimum
temperature to be considered thermal ablation, within the first half a minute and 80oC by
3 minutes [32]. Samples frozen with liquid nitrogen were assumed to surpass the
therapeutic temperature threshold. Given the extensive prior studies showing no
temperature change with IRE at the prescribed dosages or histotripsy no temperature
measurements were done for these therapies [17, 33-35]. For experimental sham,
untreated control samples of cells were prepared and handled identically to treated
samples, but instead of receiving treatment remained on wet ice during the treatment of
other cells.
Figure 8-2 Treatment flow and dosages for various ablation modalities (A). Histotripsy experiments on cell suspensions were conducted using a 1Mhz transducer (B).
309
For IRE treatments, cells were suspended in a sucrose solution, described
previously [36] to improve the quality of electrical transduction was adjusted to use a
base of PBS to mitigate differences in treatment groups, and placed into 4 mm cuvettes.
A generator (BTX ECM 830, Harvard Apparatus, Holliston, MA) was used to apply 100
pulses with widths of 100 s at a frequency of 1 Hz and an electric field of either 500
V/cm or 2000 V/cm. Cells were then cultured for 24 hours before supernatant collection
to allow for the controlled cell death of IRE to take place [15]. Additional controls for IRE
samples to accommodate for the extra downstream processing. In these cases, the
media that was used to culture the IRE samples over night was also ran through BCA
and nanodrop. The resulting values from the controls were subtracted from the IRE
samples to accommodate for the excess signaling caused by the media in culturing.
Viability for all modalities was determined shortly after treatment with standard Trypan
Blue counting with a 1:40 dilution due to the high concentration of cells, and calculated
as a percentage of cells remaining viable after treatment. Post-ablation samples were
centrifuged at 1000xg for 5 minutes and supernatants were collected. Supernatants for
cryoablation, thermal ablation, and histotripsy were collected immediately after
treatment and IRE 24 hours after incubation at 37oC.
8.4.4 In Vivo Histotripsy Treatment
Table 8-1 Subject Numbers Per Experimental Group
Immediate Group
Acute Group Chronic Group
Flow Cytometry
No Treatment No Tumor - 3 3 -
Tumor 3 7 7 4/time point
Histotripsy Treated Tumors 3 7 7 4/time point
310
Mice were treated when the average tumor diameter of the cohort was
approximately 0.6cm, to ensure that the smallest tumors were large enough for
targeting (Fig.1B). Animals were euthanized 24 hours after treatment, referred to as the
acute group, (n=7 treated, n=7 untreated controls, and n=3 tumor-free untreated
controls) or at survival time points, referred to as the chronic group (n=7 treated, n=7
untreated controls, and n=3 tumor-free untreated controls) (Table 1). Euthanasia of
chronic group animals, the point determined as their general survival, was determined
as either 1) when clinical health evaluations found deleterious symptoms including
hunching, irregular respiratory rate and rhythm, decreased alertness and socialization,
or poor extremity utilization or 2) when the tumor diameter exceeded 1.4cm. At
necropsy, the immediate group (n=3) mice that were treated and untreated (Table 1)
were euthanized and tumors were formalin fixed for histopathology to determine efficacy
of treatment. For the acute and chronic group mice, serum was collected and the
tumors were sectioned with a portion formalin fixed for histopathology and another
portion flash frozen for mRNA analysis (Fig.1B).
Prior to each treatment, each mouse had fur removed over the tumors using the
depilatory cream Nair (Naircare, Ewing, NJ). Mice were anesthetized with vaporized 2-
4% isoflurane with an oxygen flow rate of 1.5 L/min. The mice were then placed on the
subject stage with their tumor submerged in the degassed water in the subject stage’s
window. The tumor was located using the ultrasound imaging probe that was coaxially
aligned to the histotripsy transducer, and then targeted using an automated volumetric
ablation algorithm which controlled the treatment following manual targeting.
311
For each tumor, a 3D ellipsoidal volume was targeted using conservative margins of
approximately 0.5-2mm from the skin and underlying tissues (muscle, intestines, etc).
Since our automated treatment is a perfect ellipsoid and tumors are not, there was
some small amount of variation between subjects. Using these margins, we intentionally
targeted a partial ablation of approximately 60-75% of the tumor volume. This volume
consisted of multiple, concentric 2D ellipical slices. Each treatment slice was separated
0.75mm apart. Within each slice, the area was populated with grid points separated
0.75mm apart in the transverse direction and 1.25 mm apart in the axial direction
(Fig.1C). At each treatment point, histotripsy was applied at a pulse repetition frequency
(PRF) of 250Hz and a dwell time of 1 second, consequently sending 250 pulses to each
point. For each slice, the automated treatment first generated a histotripsy bubble cloud
at the center point of the slice and then scanned in a raster pattern to cover one half of
the slice area. The transducer was then returned to the center of the ellipse and
scanned in a raster pattern to cover the other half of the slice. Once a slice was
scanned through completely, the system proceeds to the next slice, which is repeated
until the entire ellipsoidal volume was treated. Throughout treatment, ultrasound
guidance confirmed the location of the bubble cloud for the duration of the volumetric
ablation (Fig.1D). After treatment, the tumor volume was again imaged with ultrasound
imaging in order to assess for tissue ablation.
312
8.4.5 Determining DNA Release and Quality
Each collected sample for each treatment group (n=5 for the no treatment group,
n=9-11/treatment group) was analyzed with a nanodrop drop, and the DNA
concentrations (ng/μL) and the 260/280 absorbance ratios were recorded. Samples
were then run on an ethidium bromide gel to visualize DNA strand sizes present in the
samples utilizing a 100bp HyperLadder (Bioline) and following standard protocol.
8.4.6 Determining Protein Release and Quality
The protein released was quantified using a BCA assay (Thermo Scientific) following
the manufacturer's protocol. Western blots were run using pre-made gels (Thermo
Scientific) following standard protocols with 5ug of protein/sample and transferred with
the iBlot 2 Dry Blotting System (Thermo Scientific). HA antigen release was determined
using an HA antibody (Cell Signaling) as per the manufacturer’s protocols. The protein’s
band area was quantified using iBright Analysis Software (n=5/treatment group).
8.4.7 Histopathology
Tissues were harvested from animals and fixed in 10% formalin. Paraffin-embedded
formalin-fixed tissues were stained with hematoxylin and eosin (H&E) following standard
protocols. Evaluations were performed by trained individuals and independently verified
by a blinded, board-certified veterinary pathologist (S.C.O.).
313
8.4.8 Profiling Gene Expression and Pathway Analysis
Using tumors flash-frozen from in vivo experiments, total RNA isolated from tumor
samples using the RNeasy Mini Kit (250) (Qiagen), where the manufacturer’s standard
protocol was followed. RNA levels were quantified with a nanodrop, converted to cDNA,
and were then pooled (n=7/tumor group and n=3/control group). Pooled cDNA was
placed into the commercially available pathway-focused array “Cancer Inflammation
and Immunity Crosstalk” (SuperArrayTM platform; Qiagen) following the manufacture’s
protocol (1/pooled groups). Fold change was determined using standard ΔΔCT
calculations. Gene expression was analyzed using IPA (integrated pathway analysis,
Qiagen) software to model changes in complex pathways [36].
Table 8-2 Flow Cytometry Immune Cell Markers
IMMUNE CELL TYPE IDENTIFYING MARKERS
Macrophages (MO) CD45+ CD11c+ F4/80+ Ly6C-
Inflammatory Dendritic Cells (iDCs) CD45+ CD11c+ F4/80- Ly6C+
Classic Dendritic Cells (cDCs) CD45+ CD11c+ F4/80- Ly6C-
Monocytic Myeloid Derived Suppressor Cells (M-MDSC) CD45+ CD11c- Ly6C+ Ly6G-
Granulocytes CD45+ CD11c- Ly6C- Ly6G+
CD8+ T cells CD45+ CD3+ CD4- CD8+
CD4+ T cells CD45+ CD3+ CD4+ CD8-
T Regulatory Cells (Treg cells) CD45+ CD3+ CD4+ CD8- FoxP3+
T Helper Cells (Th cells) CD45+ CD3+ CD4+ CD8- FoxP3-
314
8.4.9 Flow Cytometry Panel Staining
For immune cell profiling, additional mice were injected with tumors and taken down
1, 7, and 14 days after treatment (Fig.1B) with an n=4 per treated and untreated control
group at each time point (Table 1).
After removal of the tumor, avoiding skin and fur, the tumor was placed into 8mL of
cold RPMI. Even though Pan02 tumors have microvasculature,[37] because they are
not very bloody due to total blood removal via heart stick, we did not perfuse the tissue
before harvest or processing. The tissue was then mechanically digested and strained
into a 50mL conical tube. After centrifuging at 300xg for 10 minutes at 4℃, the
supernatant was discarded and the pellet resuspended in 10mL of RPMI and was
plated onto a 96-well V-bottom plate at a density of 1.0x106 cells/well. A 1:200 dilution
of FACS buffer, sterile PBS with 2% FBS and 0.1% sodium azide, and anti-CD16/32
was added to the plate at a concentration of 50µL /well. Antibodies were diluted with
FACS buffer and added directly to the wells. The following antibodies were used: anti-
CD8 SuperBright 645, anti-CD45 SuperBright 645, anti-CD11c APC, anti-CD45 PE,
anti-F4/80 FITC, anti-CD4 PE.Cy5, anti-CD8 A488, anti-CD3 APC, anti-Ly6C APC Cy5,
anti-Ly6G PE, and anti-FOXP3 PerCP Cy. Cells were washed with PBS and evaluated
with FACS (BD Biosciences). Gating for specific immune cell populations is listed in
Table 2. It should be noted that the population ‘granulocytes’ is likely to contain both
granulocytic myeloid derived suppresser cells and neutrophils [38, 39]. Additionally, by
gating macrophages as both CD11c and F4/80 we are most likely focusing on a
315
subpopulation of macrophages, more extensive panels would be necessary to confirm
the ratio of subpopulations.
8.4.10 HMGB1 Serum ELISA
Serum levels of high mobility group protein box 1 (HMGB1) in acute and survival
group mice were determined utilizing an ELISA assay kit (ABclonal) following the
manufactures suggested protocol.
8.4.11 Statistics
Data were analyzed using GraphPad Prism, version 8. Statistical significance was
defined as p ≤ 0.05, where values were not significant but p<0.20 the value is noted in
the text. All data are represented as the mean ± SEM. A Student's two-tailed t-test was
used when comparing two experimental groups. When many t-tests were performed for
one graph, group letter designations were used. Lowercase letters on top of bars
indicate significance; bars with the sample letter designation are not significant while
those that do not share a letter are significant (p<0.05). Multiple comparisons were done
using one-way or two-way ANOVA where appropriate, and then followed by Tukey post-
test for multiple pairwise examinations.
316
8.5 Results
8.5.1 Different Ablation Modalities Show Differential Release of Damage Associated
Molecular Patterns (DAMPs) and Potential Antigens
Figure 8-3 Cell suspension ablations resulted in partial and full ablations (A). DNA release was quantified (B) and visualized (C). Protein release was quantified (D) and the relative release of the antigen HA was quantified from western blot bands (E). Lowercase letters on top of bars indicate significance; bars with the sample letter designation are not significant while those that do not share a letter are significant (p<0.05).
Pan02-HA cells were treated with thermal ablation, cryoablation, IRE, and histotripsy
at full (f-) and partial (p-) doses, and were chosen based on previous literature (Fig.2A)
[16]. Treatments that lead to less than 10% viability in all samples were considered fully
ablated and those that were greater than 30% in all samples were considered partially
317
ablated (Fig.3A). All samples’ lysates were analyzed for peptide and DNA nucleotide
release (n=5 for no treatment, n=9-11 for treatment groups).
Extracellular DNA is a DAMP and often correlates with extracellular nuclear proteins,
such as HMGB1, that also acts as robust damage signaling. Here, we observed
increased levels of DNA for all treatments compared to untreated samples (Fig.3B).
DNA release was evaluated by nucleotide quantification, which showed f-IRE (470.73+/-
283.34 ng/ml) to have a significantly (p<0.05) higher concentration than most other
modalities, excluding f-cryoablation (262.49+/-159.00 ng/ml) and f-histotripsy (315.71+/-
175.28 ng/ml). P-thermal (177.33+/-113.15 ng/ml), f-thermal (170.48+/-72.20 ng/ml), p-
IRE (179.82+/-141.98 ng/ml), and p-histotripsy (165.03+83.09 ng/ml) were all similar to
each other. Only f-IRE and f-histotripsy were significantly greater than the untreated
control samples (20.35+/-10.11 ng/ml). Gel electrophoresis showed that cryoablation
and both dosages of histotripsy left large segments of detectable DNA, while untreated,
IRE and thermally ablated samples produced no visible bands (Fig.3C).
Peptide release, which correlates with potential antigens, was observed to be
released significantly in non-thermal ablation modalities (Fig.3D). There was no
significant difference between p-thermal (383.8+/-345.4 ug/ml) or f-thermal (443.3+/-
255.8 ug/ml) ablation’s peptide release from samples that were not treated (194.5+/-
50.7 ug/ml). On the other hand, f-IRE (2113+/-409.1 ug/ml) and f-histotripsy (2208+/-
751.8 ug/ml) were significantly (p<0.05) higher levels of release compared to all other
modalities, except for p-IRE (1869+/-431.3 ug/ml). F-cryoablation (1512+/-415.4 ug/ml)
and p-histotripsy (1300+/-490.0 ug/ml) were significantly different from the low
(significance group “a”) and high release clusters (significance group “c”).
318
Release of HA was confirmed on western blot for all treated samples, regardless of
ablation treatment (Sup Fig.1). The average area of the various treatments HA bands
was not significantly different between any therapies compared to each other nor the
untreated samples (297.6+/-39.3 units) (Fig.3E). The highest detection of HA was found
in p-thermal treated samples (367.8+/-24.25 units), followed by p-histotripsy (344.3+/-
38.5 units) and f-histotripsy (340.0+/-35.2 units). F-cryoablation (332.0+/-56.4 units), f-
thermal (320.3+/-33.1 units), p-IRE (322.0+/-26.4 units), and f-IRE (317.2+/-24.2 units)
were all found to have larger bands than no treatment and smaller than p-thermal, p-
histotripsy, and f-histotripsy.
8.5.2 Histotripsy is an Effective Tumor Ablation Modality in the Subcutaneous Pan02
Model
The schematic shows the custom histotripsy rig and automated ablation procedure
that was used for in vivo treatments (Fig.1). Co-axial ultrasound imaging confirmed the
presence of a bubble cloud within the tumors during treatment (Fig.1E). Mice were
treated when tumors reached 0.6cm in diameter on average and harvested in groups as
depicted (Fig.1B). Ablation of tumors was confirmed with ultrasound, with increased
hypoechoic regions, and histopathology, where a partial ablation of tumors with viable
tumor tissue on margins and in islands within the ablation zone was observed (Fig.4).
On ultrasound images, the center of the treated region had a more notable decrease in
ultrasound reflection, while the dermal margin maintained a comparable hyperechocity
after treatment compared to before (Fig.4A-B). This pattern was also noted on
319
histopathology. The center of untreated tumors has a characteristic necrotic core
(Fig.4C), and the treated tumors appeared to have a larger region of cell death that
extends nearly to the margins of the tumor (Fig.4D).
Figure 8-4 Ultrasound images of tumor before (A) and after (B) treatment, exhibiting more hypoechoic central region post-histotripsy. Orange shapes on ultrasound indicate location of tumors. Green circles indicate location of bubble cloud determined in water prior to treatment, and were utilized for treatment planning. Histology to tumors without (C) and with (D) treatment show decreased cellular detail after treatment. Dotted black line outlines the necrotic core of the tissue. Scale bar on H&E images = 500 um.
320
Figure 8-5 After treatment of tumors on day 8, as indicated by the black arrow, the reduction of tumor volume was indicated by caliper measurements (A). Changes health observed in tumor-progression free survival (B) and general survival (C).
Calculated tumor diameters decreased for a few days after the partial ablation with
histotripsy and maintained size for two weeks before resuming tumor growth (Fig.5A).
The average tumor size of untreated tumors continued to increase in size at a relatively
steady rate (Fig.5A). The greatest difference between the treated and untreated groups
was reached on day 15, 8 days post-treatment when the treated tumors were 43% of
the size of the untreated tumors on average. Compared to the untreated mice,
histotripsy treatment increased tumor-progression-free survival by 19 days from 10 days
321
to 29 days post-treatment (Fig.5B) and general survival by 8 days from 43 days to 51
days post-treatment (Fig.5C).
8.5.3 Histotripsy Ablation Results in Increased Acute Cellular Immune Response
Figure 8-6 Analysis of mRNA expression in treated tumors showed regulation of immune pathways (A). Regulated pathways interact with each other and are up regulated in the acute group and downregulated in the chronic group (B). Serum HMBG-1 levels (C) correlate with the mRNA upregulation of HMGB-1 associated pathways.
322
These tumors even without treatment tend to develop a central area of necrosis as
they progress. These necrotic cores are often characterized by accumulations of cell
debris with variable numbers of infiltrating degenerate and nondegenerate neutrophils.
Otherwise, appreciable numbers of immune cells are relatively absent microscopically
throughout the tumor tissue (Fig.6A). Treated tumors also exhibited a core of cell death
following ablation with variable infiltration by predominantly degenerate neutrophils.
However, in addition to cellular debris, these cores also often contain ghost cells and
acute hemorrhage (Fig.6B). In both treated and untreated tumors, small to moderate
numbers of neutrophils, macrophages, lymphocytes, and/or plasma ells are present at
the tumor periphery. There is no appreciable difference between the two (Fig.6C-D).
Despite the lack of appreciable differences in immune cell populations microscopically,
we investigated immune cell signaling through gene expression. Super array rtPCR
showed many genes associated with cancer inflammation and immunity crosstalk were
significantly regulated after histotripsy treatment (Sup Table 1). IPA analysis comparing
treated animals to untreated in acute and survival groups found multiple canonical
immune pathways regulated by histotripsy treatment (Fig.6A). As a trend, the pro-
inflammatory pathways are upregulated 24 hours treatment, but the majority of these
pathways become downregulated and were replaced with more anti-inflammatory
pathways at survival points. Many of the upregulated pathways (HMGB1, NF-κB, IL-6,
and TLR signaling pathways) are interconnected and are self-regulated to decrease in
function over time (Fig.6B). The schematic in Figure 5 illustrates the interactions and
mechanisms identified by IPA that are upregulated in the acute treatment group and
downregulated in the survival group. Some of these proteins and pathways were not
323
directly analyzed, but are predicted to be modulated based upon molecules that are
upstream and downstream by IPA. Together, these data indicate a significant
upregulation of pathways associated with the activation of the innate immune system
associated with DAMP signaling.
HMGB1 signaling was identified by IPA and is a potent DAMP. To verify this aspect
of our pathway analysis, we evaluated the protein levels of HMGB1 in the serum with
and without histotripsy treatment. Consistent with the IPA results, HMGB1 levels were
increased in the sera of treated mice at the acute time point compared to untreated and
control animals, while both treated and untreated mice saw a significant (p<0.05)
decrease in serum HMGB1 levels at survival endpoints (Fig.6C). Although there was a
notable increase, due to the significantly higher variance (p=0.0032), there was no
significance in the average serum HMGB1 level in the treated acute group compared to
the untreated and control groups.
8.5.4 Histotripsy Alters Immune Cell Composition in the Tumor Microenvironment
To quantify changes in immune cell populations within tumors over time after
histotripsy treatment, tumors were collected 24 hours, 7 days, and 14 days after
treatment for flow cytometry. Although there were no significant changes at the 24-hour
time point, there was an appreciable decrease in inflammatory Dendritic cells (iDCs,
p=0.19), classical Dendritic cells (cDCs, p=0.18), granulocytes, Th cells, CD4+ T cells,
and CD8+ T cell populations after treatment (Fig.6-7). The relative reduction of iDCs
(p=0.13) and cDCs (p=0.07) continued to day 7 along with a decrease in M-MDSCs, at
324
this point the remaining cells that appeared reduced at 24 hours were more comparable
to the untreated (Fig.7). At 14 days after treatment, macrophages and Treg cells were
found to be significantly reduced while cDCs were found to be significantly increased
and neutrophils (p=0.19) notably increased in treated tumors compared to the untreated
(Fig.7-8). While there we no changes in the ratios of CD4+/CD8+ T cells in the treated
tumors compared to the untreated, there were significant increases in CD4+ and a
notable increase in CD8+ T cells within the treated tumors at 7 days after treatment
(Fig.8).
Figure 8-7 Single cell suspension from treated and untreated tumors were collected at 24 hours, 7 days, and 14 days after treatment and were stained as described for flow cytometry to identify innate immune cells. Percentage of each innate immune cell population analyzed was calculated as part of the total CD45+ cells stained. Example flow cytometry plots from treated and untreated tumors at 14 days after treatment.
325
Figure 8-8 Single cell suspension from treated and untreated tumors were collected at 24 hours, 7 days, and 14 days after treatment and were stained as described for flow cytometry to identify adaptive immune cells. Percentage of each adaptive immune cell population analyzed was calculated as part of the total viable cells stained (gated as singlets). The ratio of CD4+/CD8+ T cells was calculated as a simple ratio. Example flow cytometry plots from treated and untreated tumors at 14 days after treatment.
8.6 Discussion
This study investigated the treatment of pancreatic tumors with histotripsy and
the resultant immunological responses, with a focus on the innate immune system. To
optimize our immune system assessments, we established conservative margins with
the expectation that some tumor would not be treated. This allowed cells within the
untreated margins to respond to the effects of our treatment and provided specimens for
326
subsequent analysis. For in vivo ablations, histotripsy treatment was confirmed via
bubble-cloud formation, while H&E showed complete ablation within targeted regions.
Overall, we achieved a partial ablation of all treated tumors. There was an average
reduction of 43% in the tumor diameters of the remaining tumor tissue after treatment,
comparable to tumor retardation compared to previous histotripsy subcutaneous tumor
treatments [20, 23, 24]. Based upon the fully ablated tissue found within the targeted
regions on H&E, the remaining tissue can be assumed to have been untreated based
upon the margins set. This led to tumors that were fully ablated in certain regions and
completely untreated in others. More complete ablation of tumors with histotripsy has
been achieved in de novo and in situ studies where margins include surrounding
healthy tissues, but these tumors still recurred [40-43]. In this study, the progression of
tumor growth was improved with treatment (Fig.5B). However, in line with previous
studies, tumor recurrence occurred and minimized the improvement of the general
survival for animals that received treatment (Fig.5C).
The release of DAMPs is a consistent feature of focal ablation modalities, and
the established modalities have been compared to each other in previous work [16].
Histotripsy has not been previously, directly compared to non-ultrasonic ablation
modalities. The effects of histotripsy compared to other ablation modalities for producing
immune-stimulating molecules is of interest given the response to the histotripsy
treatment of pancreatic tumors in vivo yielded an inflammatory response that is turned
off over time while showing relatively large variations of immune cell proportions within
treated tumors. Looking at the release of DNA and peptides as potential DAMPs,
327
showed that histotripsy is comparable to the non-thermal ablations cryoablation and IRE
(Fig.3).
Extracellular DNA is recognized by innate immune system receptors as a DAMP
and is not typically found in the absence of damage [44]. Additionally, the presence of
released proteins increases the probability of the immune system being recruited by
damage associated cytokines and antigens. Our hypothesized expectations for the in
vitro studies were that cryoablation and histotripsy should have similar, more intact
DNA, in larger fragments, and more protein released from cells after ablation given that
they both ablate cells through immediate lysis [17, 45]. On the other hand, no cell death
was observed in untreated cells, also as expected, and therefore had low levels of
protein and DNA. However, IRE induces a delayed cell death, defined as either
apoptosis or pyroptosis [15]. DNA fragmentation and protein release or damage are
hallmark features of programed cell death. Similarly, thermal ablation is known to
directly denature DNA and protein [16, 46]. Thus, in both of these cases, we predicted
that the DNA released following ablation would be significantly more fragmented,
including significantly smaller and undetectable fragments. Even though IRE does use a
delayed cell death mechanism, we did expect to find the higher level of detectable
proteins given that programmed cell death does not denature all of the released
proteins and previous studies that also showed that IRE can release a significant
magnitude of intact proteins [15, 16].
In addition, results from the in vitro experiments provide important comparisons
between different ablation methods and demonstrate that non-thermal ablation
approaches, including histotripsy, lead to significantly increased release of DAMPs and
328
potential antigens in comparison to thermal ablation methods. In comparing the clinical
relevance of the differential release of DAMPs and potential antigens that was observed
between the ablation modalities in vitro, it is important to note that additional in vivo
studies will be needed to further study the differences between each ablation modality.
Similarly, additional in vivo studies will be needed to further investigate the role of
treatment dose in immune system activation for each of the ablation modalities. For
instance, it is not clear whether the partial ablations generated in our in vitro studies will
be representative of a partial or incomplete ablation in a clinical setting. For the thermal
ablation, cryoablation, and IRE samples treated at partial-ablation doses in this study in
vitro, all of the cells in suspension were treated at a reduced magnitude. In contrast,
since histotripsy is a binary treatment that requires a bubble cloud to be formed once a
distinct pressure threshold has been exceeded [47-50], the histotripsy partial ablation
resulted in only a portion of the cells being exposed to a full amplitude histotripsy bubble
cloud, whereas the remaining cells received no treatment. When comparing these in
vitro results from cell suspensions to in vivo ablations, it is important to remember that
these differences in how a partial ablation versus a full ablation is acquired may result in
different responses. As a result, controlled in vivo studies comparing the effects of
different ablation modalities as well as different treatment doses on the potential
immunological benefits should be conducted. In addition to comparing the differences
between partial and complete ablation, these future studies should investigate the
potential risks of overtreatment that could potentially reduce immunological benefits. For
instance, in prior work designing IRE protocols, users have outlined specific treatment
guidelines to avoid excessive Joule heating to capitalize on the therapeutic benefits of
329
the non-thermal IRE ablation [34]. Similarly, samples over treated with thermal ablation
modalities and cryoablation face an increased rate of macromolecule breakdown and
protein denaturation compared to non-thermal [16]. Future studies are needed to
determine any effects of overtreatment with histotripsy and to determine the optimal
histotripsy treatment strategies for maximizing immunological benefits.
Given that histotripsy’s mechanism of ablation is the mechanical lysis of cells and
that in vitro we found increased release of DNA and proteins (Fig.3), the determination
that DAMP signaling pathways (Fig.5F) were activated in vivo was not surprising.
Although we did not appreciate histological changes in immune cell infiltration on a
microscopic level, gene expression analysis suggested changes in immune cell
signaling pathways which led us to assess immune cell populations via flow cytometry,
a more sensitive indicator. Here we were able to identify specific pathways modulated
by histotripsy, including HMGB1, IL-6, Interferon, TEC Kinase, JAK/STAT, and PPAR
pathway signaling. These pathways are consistent with responses to trauma or physical
damage [51-53]. As time passes after a trauma, these signaling pathways become
repressed to control the immune response. One way this is done is through the PPAR
pathway, which can inhibit the downstream production of cytokines from the HMGB1
and NF-κB pathways [54]. The activation of the PPAR pathway can also increase
healing effects by stimulating the production of VEGF [55], found in our study to be
significantly increased in treated survival group tumors (Fig.5F, Sup Table 1). For
healing, the production of VEGF is needed to reestablish healthy vasculature; however,
high levels of VEGF within a tumor have been established to be tumorigenic and
correlates with poor patient outcomes [56]. Histotripsy increasing the activation of the
330
PPAR pathway and increasing the levels of VEGF expression did not promote negative
outcomes for the animals in our study but could be targets of future adjuvants to extend
the early inflammatory tumor microenvironment.
At two weeks post-histotripsy ablation, there was a significant decrease in
macrophages (Fig.6) and Treg cells (Fig.7). Significant decreases in the expression of
cytokines associated with tumor-associated macrophages (TAMs) including IL-4, IL-10
CCL-2, CCL-22, and CXCl-12 (Sup. Table 1) and suggest that the reduction in
macrophages within the treated tumors could be indicative of a decrease in TAMs [57,
58]. A decrease in Treg cells with a potential decrease in TAMs could indicate additional
access of histotripsy immunomodulation of the tumor microenvironment to being less
anti-inflammatory. This could be a potential target for enhancement with adjuvant
therapies [57, 59]. Overall, these changes to immune cell populations should be further
analyzed in future studies to determine the full extent of changes to sub-populations,
such as changes to M1/M2/TAM macrophages instead of basic populations shown
here.
Early studies investigating the immunological effects of histotripsy compared the
effects of acoustic cavitation to acoustic heating with thermal HIFU. One study using a
subcutaneous colon adenocarcinoma mouse model showed that histotripsy is capable
of stimulating CD11c+ cells within the tumors more than thermal HIFU [26]. It has also
been shown that histotripsy of murine melanoma can better stimulate an immune
response with a decrease in metastasis in the weeks following treatment compared to
HIFU, suggesting involvement of anti-tumor lymphocytes [27]. More recent studies have
further established the pro-inflammatory local and systemic effects of histotripsy on the
331
immune response in melanoma, neuroblastoma, hepatocellular carcinoma, and renal
cell carcinoma [22-24]. This study adds to this knowledge by establishing a framework
for the immune response to histotripsy ablation of pancreatic cancer. The response
reestablished here is similar to other studies. However, given that the Pan02 tumors are
known to be poorly immunogenic [60], it was not surprising that the local changes in
immune cell populations, while significant at points (Figs.7-8), were not as prominent of
a profile shift as what has been reported in other tumors types. Using the Pan02 model,
a prior study with IRE showed that subcutaneous tumors did not have a strong of a
change in immune cell populations within the tumor as orthotopic tumors [61]. Similar to
the current work showed data that non-thermal ablation of the subcutaneous pancreatic
tumors can shift the tumor microenvironment to being more pro-inflammatory [61].
The results of this study provide evidence that histotripsy can ablate subcutaneous
pancreatic tumors and stimulate a local immune response. This study builds upon
previous studies utilizing histotripsy for other tumor types [20, 22-24], and shows a
potential immune profile for pancreatic tumors after histotripsy ablation. Overall, the
results of this work provide a baseline expectation of the response of pancreatic tumors
to histotripsy, which will help for planning future orthotopic studies.
8.7 Conclusion
This study demonstrates the feasibility of using histotripsy for pancreatic cancer
ablation and defines mechanisms associated with innate immune system activation
332
following treatment. Further work is needed to establish the mechanisms behind the
immunomodulation of the tumor microenvironment and systemic effects.
8.8 Acknowledgment
This work was supported by the Virginia Maryland College of Veterinary Medicine; The
Virginia Tech Institute for Critical Technology and Applied Sciences Center for
Engineered Health; and The National Institutes of Health. The content is solely the
responsibility of the authors and does not necessarily represent the official views of the
NIH or any other funding agency. Figures 1B and 6F were created using biorender.com.
333
8.9 References
[1] R. L. Siegel, K. D. Miller, and A. Jemal, "Cancer statistics, 2017," CA: a cancer
journal for clinicians, vol. 67, no. 1, pp. 7-30, 2017.
[2] R. L. Siegel, K. D. Miller, and A. Jemal, "Cancer statistics, 2020," CA: a cancer
journal for clinicians, vol. 70, no. 1, pp. 7-30, 2020.
[3] H. Manuel, "Pancreatic cancer," New England Journal of Medicine, vol. 362, no.
17, 2010.
[4] S. Gillen, T. Schuster, C. M. Zum Büschenfelde, H. Friess, and J. Kleeff,
"Preoperative/neoadjuvant therapy in pancreatic cancer: a systematic review and
meta-analysis of response and resection percentages," PLoS medicine, vol. 7,
no. 4, 2010.
[5] O. Kepp, A. Marabelle, L. Zitvogel, and G. Kroemer, "Oncolysis without viruses—
inducing systemic anticancer immune responses with local therapies," Nature
Reviews Clinical Oncology, pp. 1-16, 2019.
[6] C. J. Simon, D. E. Dupuy, and W. W. Mayo-Smith, "Microwave ablation:
principles and applications," Radiographics, vol. 25, no. suppl_1, pp. S69-S83,
2005.
[7] Y. Keisari, "Tumor abolition and antitumor immunostimulation by physico-
chemical tumor ablation," Front Biosci (Landmark Ed), vol. 22, pp. 310-47, 2017.
[8] R. V. Davalos, L. Mir, and B. Rubinsky, "Tissue ablation with irreversible
electroporation," Annals of biomedical engineering, vol. 33, no. 2, p. 223, 2005.
334
[9] R. C. Martin et al., "Treatment of 200 locally advanced (stage III) pancreatic
adenocarcinoma patients with irreversible electroporation: safety and efficacy,"
Annals of surgery, vol. 262, no. 3, pp. 486-494, 2015.
[10] H. J. Scheffer et al., "Ablation of locally advanced pancreatic cancer with
percutaneous irreversible electroporation: results of the phase I/II PANFIRE
study," Radiology, vol. 282, no. 2, pp. 585-597, 2017.
[11] M. P. Belfiore et al., "Percutaneous CT-guided irreversible electroporation
followed by chemotherapy as a novel neoadjuvant protocol in locally advanced
pancreatic cancer: our preliminary experience," International Journal of Surgery,
vol. 21, pp. S34-S39, 2015.
[12] H. J. Scheffer et al., "Irreversible electroporation of locally advanced pancreatic
cancer transiently alleviates immune suppression and creates a window for
antitumor T cell activation," Oncoimmunology, vol. 8, no. 11, p. 1652532, 2019.
[13] R. C. Martin et al., "Irreversible electroporation in locally advanced pancreatic
cancer: a call for standardization of energy delivery," Journal of Surgical
Oncology, vol. 114, no. 7, pp. 865-871, 2016.
[14] N. Beitel-White, R. C. G. Martin, Y. Li, R. M. Brock, I. C. Allen, and R. V.
Davalos, "Real-time prediction of patient immune cell modulation during
irreversible electroporation therapy," (in eng), Sci Rep, vol. 9, no. 1, p. 17739,
Nov 28 2019, doi: 10.1038/s41598-019-53974-w.
[15] R. M. Brock, N. Beitel-White, R. V. Davalos, and I. C. Allen, "Starting a Fire
Without Flame: The Induction of Cell Death and Inflammation in Electroporation-
Based Tumor Ablation Strategies," Frontiers in Oncology, vol. 10, 2020.
335
[16] Q. Shao et al., "Engineering T cell response to cancer antigens by choice of focal
therapeutic conditions," International Journal of Hyperthermia, vol. 36, no. 1, pp.
130-138, 2019.
[17] K. B. Bader, E. Vlaisavljevich, and A. D. Maxwell, "For whom the bubble grows:
physical principles of bubble nucleation and dynamics in histotripsy ultrasound
therapy," Ultrasound in medicine & biology, 2019.
[18] E. Vlaisavljevich, A. Maxwell, L. Mancia, E. Johnsen, C. Cain, and Z. Xu,
"Visualizing the Histotripsy Process: Bubble Cloud-Cancer Cell Interactions in a
Tissue-Mimicking Environment," (in Eng), Ultrasound Med Biol, Jul 8 2016, doi:
S0301-5629(16)30099-0 [pii] 10.1016/j.ultrasmedbio.2016.05.018.
[19] E. Vlaisavljevich et al., "Non-invasive ultrasound liver ablation using histotripsy:
Chronic study in an in vivo rodent model," Ultrasound in medicine & biology, vol.
42, no. 8, pp. 1890-1902, 2016.
[20] T. Worlikar et al., "Histotripsy for Non-Invasive Ablation of Hepatocellular
Carcinoma (HCC) Tumor in a Subcutaneous Xenograft Murine Model," in 2018
40th Annual International Conference of the IEEE Engineering in Medicine and
Biology Society (EMBC), 2018: IEEE, pp. 6064-6067.
[21] K. C. Longo et al., "Robotically Assisted Sonic Therapy (RAST) for Noninvasive
Hepatic Ablation in a Porcine Model: Mitigation of Body Wall Damage with a
Modified Pulse Sequence," Cardiovasc Intervent Radiol, vol. 42, no. 7, pp. 1016-
1023, Jul 2019, doi: 10.1007/s00270-019-02215-8.
[22] G. R. Schade, Y.-N. Wang, S. D'Andrea, J. H. Hwang, W. C. Liles, and T. D.
Khokhlova, "Boiling Histotripsy Ablation of Renal Cell Carcinoma in the Eker Rat
336
Promotes a Systemic Inflammatory Response," Ultrasound in medicine &
biology, vol. 45, no. 1, pp. 137-147, 2019.
[23] A. Eranki et al., "High-Intensity Focused Ultrasound (HIFU) Triggers Immune
Sensitization of Refractory Murine Neuroblastoma to Checkpoint Inhibitor
Therapy," Clinical Cancer Research, vol. 26, no. 5, pp. 1152-1161, 2020.
[24] S. Qu et al., "Non-thermal histotripsy tumor ablation promotes abscopal immune
responses that enhance cancer immunotherapy," Journal for ImmunoTherapy of
Cancer, vol. 8, no. 1, 2020.
[25] K. J. Pahk et al., "Boiling Histotripsy-induced Partial Mechanical Ablation
Modulates Tumour Microenvironment by Promoting Immunogenic Cell Death of
Cancers," Scientific reports, vol. 9, no. 1, pp. 1-12, 2019.
[26] Z. Hu et al., "Investigation of HIFU-induced anti-tumor immunity in a murine
tumor model," Journal of translational medicine, vol. 5, no. 1, p. 34, 2007.
[27] Y. Xing, X. Lu, E. C. Pua, and P. Zhong, "The effect of high intensity focused
ultrasound treatment on metastases in a murine melanoma model," Biochemical
and biophysical research communications, vol. 375, no. 4, pp. 645-650, 2008.
[28] M. E. Lorkowski et al., "Immunostimulatory nanoparticle incorporating two
immune agonists for the treatment of pancreatic tumors," (in eng), J Control
Release, Nov 11 2020, doi: 10.1016/j.jconrel.2020.11.014.
[29] C. C. Baniel et al., "In situ Vaccine Plus Checkpoint Blockade Induces Memory
Humoral Response," (in eng), Front Immunol, vol. 11, p. 1610, 2020, doi:
10.3389/fimmu.2020.01610.
337
[30] A. D. Maxwell, C. A. Cain, A. P. Duryea, L. Yuan, H. S. Gurm, and Z. Xu,
"Noninvasive thrombolysis using pulsed ultrasound cavitation therapy–
histotripsy," Ultrasound in medicine & biology, vol. 35, no. 12, pp. 1982-1994,
2009.
[31] B. Aarts et al., "Cryoablation and immunotherapy: an overview of evidence on its
synergy," Insights into imaging, vol. 10, no. 1, pp. 1-12, 2019.
[32] C. Brace, "Thermal tumor ablation in clinical use," (in eng), IEEE Pulse, vol. 2,
no. 5, pp. 28-38, Sep-Oct 2011, doi: 10.1109/mpul.2011.942603.
[33] M. Faroja et al., "Irreversible electroporation ablation: is all the damage
nonthermal?," Radiology, vol. 266, no. 2, pp. 462-470, 2013.
[34] R. V. Davalos, S. Bhonsle, and R. E. Neal, "Implications and considerations of
thermal effects when applying irreversible electroporation tissue ablation
therapy," The Prostate, vol. 75, no. 10, pp. 1114-1118, 2015.
[35] R. M. Brock et al., "Patient derived xenografts expand human primary pancreatic
tumor tissue availability for ex vivo irreversible electroporation testing," Frontiers
in Oncology, vol. 10, 2020.
[36] V. M. Ringel-Scaia et al., "High-frequency irreversible electroporation is an
effective tumor ablation strategy that induces immunologic cell death and
promotes systemic anti-tumor immunity," EBioMedicine, vol. 44, pp. 112-125, Jun
2019, doi: 10.1016/j.ebiom.2019.05.036.
[37] M. Kato, K. Watabe, M. Tsujii, T. Funahashi, I. Shimomura, and T. Takehara,
"Adiponectin inhibits murine pancreatic cancer growth," Digestive diseases and
sciences, vol. 59, no. 6, pp. 1192-1196, 2014.
338
[38] J. I. Youn and D. I. Gabrilovich, "The biology of myeloid‐derived suppressor cells:
the blessing and the curse of morphological and functional heterogeneity,"
European journal of immunology, vol. 40, no. 11, pp. 2969-2975, 2010.
[39] S. Rose, A. Misharin, and H. Perlman, "A novel Ly6C/Ly6G‐based strategy to
analyze the mouse splenic myeloid compartment," Cytometry Part A, vol. 81, no.
4, pp. 343-350, 2012.
[40] N. R. Styn, T. L. Hall, J. B. Fowlkes, C. A. Cain, and W. W. Roberts, "Histotripsy
of renal implanted VX-2 tumor in a rabbit model: investigation of metastases,"
Urology, vol. 80, no. 3, pp. 724-729, 2012.
[41] G. R. Schade et al., "Histotripsy focal ablation of implanted prostate tumor in an
ACE-1 canine cancer model," The Journal of urology, vol. 188, no. 5, pp. 1957-
1964, 2012.
[42] T. Worlikar et al., "Non-Invasive Orthotopic Liver Tumor Ablation Using
Histotripsy in an in Vivo Rodent Hepatocellular Carcinoma (HCC) Model,"
presented at the Society of Interventional Oncology, Boston, MA, USA, 2019.
[43] J. Vidal Jove et al., "Phase I Study of Safety and Efficacy of Hepatic Histotripsy:
Preliminary Results of First in Man Experience with Robotically-Assisted Sonic
Therapy," presented at the International Society of Therapeutic Ultrasound,
Barcelona, Spain, 2019.
[44] D. S. Pisetsky, "The origin and properties of extracellular DNA: from PAMP to
DAMP," Clinical Immunology, vol. 144, no. 1, pp. 32-40, 2012.
339
[45] J. P. Erinjeri and T. W. Clark, "Cryoablation: mechanism of action and devices,"
Journal of Vascular and Interventional Radiology, vol. 21, no. 8, pp. S187-S191,
2010.
[46] K. F. Chu and D. E. Dupuy, "Thermal ablation of tumours: biological mechanisms
and advances in therapy," Nature Reviews Cancer, vol. 14, no. 3, pp. 199-208,
2014.
[47] E. Vlaisavljevich, T. Gerhardson, T. Hall, and Z. Xu, "Effects of f-number on the
histotripsy intrinsic threshold and cavitation bubble cloud behavior," Phys Med
Biol, vol. 62, no. 4, pp. 1269-1290, Feb 21 2017, doi: 10.1088/1361-
6560/aa54c7.
[48] E. Vlaisavljevich et al., "Effects of Ultrasound Frequency and Tissue Stiffness on
the Histotripsy Intrinsic Threshold for Cavitation," Ultrasound Med Biol, 2015.
[49] E. Vlaisavljevich, A. Maxwell, M. Warnez, E. Johnsen, C. A. Cain, and Z. Xu,
"Histotripsy-induced cavitation cloud initiation thresholds in tissues of different
mechanical properties," IEEE Trans Ultrason Ferroelectr Freq Control, vol. 61,
no. 2, pp. 341-52, Feb 2014, doi: 10.1109/TUFFC.2014.6722618.
[50] A. D. Maxwell, C. A. Cain, T. L. Hall, J. B. Fowlkes, and Z. Xu, "Probability of
cavitation for single ultrasound pulses applied to tissues and tissue-mimicking
materials," Ultrasound in medicine & biology, vol. 39, no. 3, pp. 449-465, 2013.
[51] D. S. Pisetsky, "Cell death in the pathogenesis of immune-mediated diseases:
the role of HMGB1 and DAMP-PAMP complexes," Swiss medical weekly, vol.
141, p. w13256, 2011.
340
[52] E. Venereau, M. Schiraldi, M. Uguccioni, and M. E. Bianchi, "HMGB1 and
leukocyte migration during trauma and sterile inflammation," Molecular
immunology, vol. 55, no. 1, pp. 76-82, 2013.
[53] N. Haque et al., "Role of the CXCR4-SDF1-HMGB1 pathway in the directional
migration of cells and regeneration of affected organs," World Journal of Stem
Cells, vol. 12, no. 9, p. 938, 2020.
[54] S. Ying, X. Xiao, T. Chen, and J. Lou, "PPAR ligands function as suppressors
that target biological actions of HMGB1," PPAR research, vol. 2016, 2016.
[55] L. Piqueras et al., "Activation of PPARβ/δ induces endothelial cell proliferation
and angiogenesis," Arteriosclerosis, thrombosis, and vascular biology, vol. 27,
no. 1, pp. 63-69, 2007.
[56] A. Lapeyre-Prost et al., "Immunomodulatory activity of VEGF in cancer," in
International review of cell and molecular biology, vol. 330: Elsevier, 2017, pp.
295-342.
[57] J. Zhou, Z. Tang, S. Gao, C. Li, Y. Feng, and X. Zhou, "Tumor-Associated
Macrophages: Recent Insights and Therapies," Frontiers in Oncology, vol. 10, p.
188, 2020.
[58] G. J. Szebeni, C. Vizler, K. Kitajka, and L. G. Puskas, "Inflammation and cancer:
extra-and intracellular determinants of tumor-associated macrophages as tumor
promoters," Mediators of inflammation, vol. 2017, 2017.
[59] B. Chaudhary and E. Elkord, "Regulatory T cells in the tumor microenvironment
and cancer progression: role and therapeutic targeting," Vaccines, vol. 4, no. 3,
p. 28, 2016.
341
[60] A. Kumar, K. Chamoto, P. S. Chowdhury, and T. Honjo, "Tumors attenuating the
mitochondrial activity in T cells escape from PD-1 blockade therapy," ELife, vol.
9, p. e52330, 2020.
[61] C. He, X. Huang, Y. Zhang, X. Lin, and S. Li, "T‐cell activation and immune
memory enhancement induced by irreversible electroporation in pancreatic
cancer," Clinical and Translational Medicine, vol. 10, no. 2, p. e39, 2020.
342
8.10 Supplemental Table and Figure Legends
Figure 8-9 Supplemental Figure 1: Whole western blot images analyzing HA antigen released in ablated cell supernatants.
343
Table 8-3 Supplemental Table 1: Genes analyzed from SuperArry rtPCR and calculated fold regulation.
344
Chapter 9. Conclusion and Future Directions
345
9.1 Introduction
This dissertation consists of three parts. Part One of this dissertation (Chapters 2- 3)
focuses on determining the safety and dosages for using histotripsy to treat stronger
hepatopancreatobiliary tumors using excised tissues and patient derived xenograft
(PDX) mice. Part Two (Chapters 4-6) moves onto more complex models, using large
animals and veterinary patients, to establish application protocols for additional tumor
types. Part Three (Chapters 7-8) shifts to utilizing immunocompetent mouse models to
investigate the immunological effects of histotripsy in pancreatic adenocarcinoma.
9.2 How can we safely and efficacious ablate more difficult tumors?
In Chapter 2, the role of histotripsy dose in the susceptibility of various liver
tumors is investigated. Specifically, it was determined that while using a 700KHz
histotripsy transducer with a PRF of 500Hz, hepatocellular carcinoma tumors can be
fully ablated with an applied dose 500 pulses/point (Fig.2-5). However, when looking at
cholangiocarcinoma (CC) tumors, known to be highly fibrotic and have a higher Young’s
modulus than most other liver malignancies [REF], even at a dosage of 4000
pulses/point there was about 25% cell viability within the targeted area (Fig.2-7E).
Even though this ex vivo study was limited by sample numbers, due to the rarity of
CC, a patient-derived xenograft (PDX) tumor model of CC was utilized to determine if it
could be safe and practical to achieve a significant ablation with histotripsy. In two
346
different PDX cohorts, with tumors derived from different patients, it was found that
using a small, 8-element 1MHz transducer at a PRF of 500Hz, that a complete ablation
of targeted tumor was achieved at 500 pulses/point (Fig.2-3K and Fig.3-2B). Knowing
that it was feasible to ablate the PDX CC tumors with a reasonable dose, Chapter 3
focused on determining whether ablation of the full tumor volume is safe. When
applying a 1-2mm positive margin to ensure the application of histotripsy to the entire
CC tumor, the off-target damage was deemed too great to justify (Fig.3-3E-F).
However, when applying a 0mm margin, therefore attempting to target 100% of the
tumor and no margin, the ablation safety was significantly higher, leading to the only
adverse events seen being mild bruising immediately after treatment and a small scab
within the first week (Fig.3-3D). Within 2.5 weeks of treatment, as the ablated
homogenate was resorbed, the decrease in tumor diameter was found to reach a
maximum of 73%, when compared to the untreated tumors, including the temporary
disappearance of 3 of the 6 treated tumors (Fig.3-4). Future work for CC tumor ablation
needs to determine how to utilize an effective ablation dose on the tumor to achieve a
full ablation without causing significant damage to off target tissues.
9.3 Can we use a large animal model to develop histotripsy for acoustically
difficult to target tumor locations?
As discussed in Chapter 4, testing histotripsy devices in rodents requires smaller,
custom made therapeutic systems (Fig.4-1), while utilizing large animal models and
veterinary patients allows for the testing of human scale systems (Fig.4-2). Pilot studies
347
of canine soft tissue sarcoma and osteosarcoma demonstrate the feasibility of utilizing
these de novo tumor carrying animals to determine therapeutic protocols for histotripsy
(Fig.4-7,8).
While there are many benefits to using veterinary patients, one drawback is the
lack of easy replication of exact tumor size, location, and severity that is offered by
tumor models. Chapters 5 and Appendix A lay the foundation for our newly
established porcine tumor model. By using CRISPR/Cas9 genetically modified pigs that
lack function RAG2 and IL-2RG proteins (Fig.6-1), which led the pigs to be severely
immunocompromised (Fig.6-2), a range of established human and murine cell lines can
be engrafted for tumor generation (Fig.4-4, 5-3, A-2). Focusing investigations on
pancreatic cancer, the engraftment of the human pancreatic adenocarcinoma cell line,
Panc01, showed luminal structures (Fig.5-5) and expression of the cytokeratin protein
CK-19 (Fig.5-6) that is consistent with human pancreatic tumors. While this study
established a porcine model for pancreatic cancer, before using these animals it is
necessary to establish a safe and effective protocol for targeting the pancreas with
histotripsy.
Chapter 6 investigates the potential of using histotripsy to target the porcine
pancreas. The goal of this project was to determine the safety of the treatment as well
as establishing the least invasive protocol for using histotripsy. The first pilot study
demonstrated that with minimal intestinal gas reduction (i.e. fasting for 12 hours)
resulted in poor acoustic visualization of the pancreas (Fig.6-3B,4) and mild damage to
the intestines (Fig.6-4). For subsequent studies, the animals were fed a custard that
contained laxative and simethicone, the combination of which allowed for the clear
348
visualization of the pancreas (Fig.6-3C) and histotripsy ablation (Fig.6-5). Using this
protocol, 4 animals were survived one week with no negative side effects noted in
behavior, diet, blood work (Fig.6-7), CT imaging (Fig.6-8), or necropsy (Fig.6-6,9). This
study established a protocol for safely targeting the pancreas for ablation with
histotripsy. Future work needs to determine the feasibility of this protocol for targeting
tumors within the pancreas.
9.4 How do different tumor types immunologically respond to histotripsy?
Chapter 7 establishes the foundation of histotripsy’s role in immunomodulation.
This literature review establishes the pre-clinical immune effects caused by the
mechanical ablation of tumors with histotripsy. This includes production of damage
associated molecular patterns, release of immune stimulating cytokines, changes in
local immune cell populations to those that are more pro-inflammatory and anti-tumor,
and increases in systemic immunity (Fig.7-5). However, this review also serves to
highlight future work that is still needed, including investigations into repolarization of
pro-tumor immune cells, synergistic effects of check-point inhibitors and other
chemotherapeutics, and the full extent of systemic, anti-body mediated immunity
stimulated against the tumor. In sum, these studies suggest that there is a reproducible,
consistent, and perhaps even tunable immune effect generated by histotripsy modalities
that is consistent across multiple tumor types. This review also highlights that all of this
work has been done utilizing small animal models and only across a few tumor types. In
order to fully understand the scope of the immunological effects of histotripsy, more
349
work needs to be done in additional tumor types, large animals, and human clinical
trials. Additionally, future work should investigate how histotripsy, a non-thermal and
non-invasive, ablation’s effects compare to other, established ablation modalities.
Following this, Chapter 8 focused on determining the potential immune effects of
histotripsy on pancreatic cancer using the murine Pan02 model. To study the effects of
histotripsy on pancreatic cancer, we utilized an in vitro model of pancreatic
adenocarcinoma and compared the release of neoantigens following histotripsy
treatment to other ablation modalities (Fig.8-2). Histotripsy was found to release
immune-stimulating molecules at magnitudes similar to other non-thermal ablation
modalities and superior to thermal ablation modalities, which corresponded to increased
innate immune system activation in vivo (Fig.8-3). In subsequent in vivo studies, murine
Pan02 tumors were treated with histotripsy, and observed significantly increased
progression-free and overall survival (Fig.8-5), with increased activation of the innate
immune system 24 hours post-treatment (Fig.8-6) and decreased tumor-associated
immune cell populations within 14 days of treatment (Fig.8-7,8). This study
demonstrates the feasibility of using histotripsy for pancreatic cancer ablation and
provides mechanistic insight into the initial innate immune system activation following
treatment. Further work is needed to establish the mechanisms behind the
immunomodulation of the tumor microenvironment and systemic effects.
350
9.5 Overall Conclusion
The results of this dissertation provide insight into establishing protocols for treating
new types of tumors with histotripsy and immunological effects that lay groundwork for
improving future co-therapeutic planning. Future work will aim to translate histotripsy
into clinical applications and determining co-therapies that can improve local and
systemic tumor burden.
351
Appendix A. Employing Novel Porcine Models of Subcutaneous Pancreatic Cancer to Evaluate Oncological Therapies
Alissa Hendricks-Wenger, Margaret A Nagai-Singer, Kyungjun Uh, Eli Vlaisavljevich, Kiho Lee, Irving C Allen
This chapter is excerpted from a textbook chapter accepted to “Bioengineering
Technology,” Humana Press.
352
A.1 Abstract
Immunocompromised mice are commonly utilized to study pancreatic cancer and other
malignancies. The ability to xenograft tumors in either subcutaneous or orthotopic
locations provides a robust model to study diverse biological features of human
malignancies. However, there is a dire need for large animal models that better
recapitulate human anatomy in terms of size and physiology. These models will be
critical for biomedical device development, surgical optimization, and drug discovery.
Here, we describe the generation and application of immunocompromised pigs lacking
RAG2 and IL2RG as a novel model for human xenograft studies. These SCID-like pigs
closely resemble NOD scid gamma mice and are receptive to human tumor tissue, cell
lines, and organoid xenografts. However, due to their immunocompromised nature,
these immunocompromised animals require housing and maintenance under germfree
conditions. In this protocol, we describe the use of these pigs in a subcutaneous tumor
injection study with human PANC1 cells. The tumors demonstrate a steady, linear
growth curve, reaching 1.0 cm within 30 days post-injection. The model described here
is focused on subcutaneous injections behind the ear. However, it is readily adaptable
for other locations and additional human cell types.
A.2 Keywords
Immunocompromised, Porcine, Cancer, Pancreatic, Ultrasound, Therapy
353
A.3 Introduction
Pancreatic cancer is expected to become the second leading cause of cancer
related deaths by 2020, with patient survival rates below 9% due to late diagnosis and
poor treatment options.[1] The current armamentarium for treating pancreatic cancer is
rather limited, but still includes standard oncological care: surgery, chemotherapy,
radiation, immunotherapy, and ablation therapy. Unfortunately, the most effective
treatment option is surgical resection of the tumor, which is not a viable strategy for
patients that have advanced disease or co-morbidities.[2] The frontline chemotherapy,
gemcitabine, has been one of the most significant improvements to standard of care for
pancreatic cancer, but only improved the one-year survival from 2% to 18%.[2,3] The
scarcity of treatment options is due to late diagnosis, complicated stromal tissue, and
lack of appropriate models for studying human disease predictably in pre-clinical
studies.
In recent years, advancements in cancer therapeutics, such as minimally
invasive ablation modalities, have shown promise in pre-clinical trials and early clinical
work. For example, irreversible electroporation (IRE) ablates tumors by generating
short, high-amplitude electric fields, preferentially killing tumor cells in the ablation zone,
while leaving healthy tissue relatively intact.[4-6] Human trials have shown promise for
IRE in ablating pancreatic tumors, with patients having significant increases in overall
survival and minimal adverse events.[7-11] Focused Ultrasound therapies have also
been developed for non-invasive tumor ablation, including high intensity focused
ultrasound (HIFU) and histotripsy. HIFU has been developed as a thermal ablation
modality and has been found to improve patient outcomes in pre-clinical and clinical
354
trials.[12,13]Histotripsy is a non-invasive focused ultrasound ablation technology that
uses cavitation at a focal point to ablate the targeted tissue into an acellular liquid
homogenate.[14] With the precision and tissue selectivity seen in pre-clinical trials to
date, there is hope that histotripsy will be able to overcome the negative side effects
typically reported with other more invasive ablation modalities.[15-17] However, with
IRE, histotripsy, and other emerging technologies, there are still many questions that
need to be answered about safety and efficacy that are difficult to address using
currently available animal models.
In order to better recapitulate human disease, traditional approaches have
focused on using genetically modified mouse lines.[18,19] While mice have proven to
be valuable for the study of basic cancer biology, they often fail to seamlessly translate
therapeutic strategies to human disease due to their size, differences in immune
system, lifespan, telomerase activity, genome, and other basic biological functions.
Thus, there is a need for an animal model that more predictably recapitulates human
anatomy and physiology. This need can be addressed utilizing pigs. As a model
organism pigs are similar to humans in size, genome, metabolism, telomerase activity,
and organ micro-structures.[20,21] Genetically modified pig models have included
animals with APC and TP53 mutations that developed colorectal polyps and osteogenic
tumors, respectively, similar in pathogenesis to human malignancies.[22,23] In order to
increase the breadth of tumorigenic sites, the “Oncopig” was developed with KRAS and
TP53 mutations. When stimulated by Cre recombinase activation, these pigs develop
tumor nodules at the site of Cre injection. [24,20] However, while these models have
been useful for specific studies, they are associated with significant limitations that have
355
limited their use for many applications. Specifically, these models do not fully
recapitulate human disease, are not reproducible on a short time scale, and lack the
complete control of features such as growth dynamics and tumor location that is
available in well-established murine models.
In order to address this specific niche, we have generated pigs with severe
combined immunodeficiency (SCID)-like disease that can be used to reliably grow
tumors and are receptive to engraftment with human cells. These pigs are designed to
be engrafted with a diverse range of human tumor cell lines and tissues either
orthotopically or subcutaneously. The tumors can be implanted in highly controlled sites
within each tissue (i.e. in close proximity to critical structures, such as blood vessels or
ducts), and the growth curve for xenografts can be highly standardized. There are two
established means to achieve SCID-like pigs: naturally occurring or TALEN-mediated
mutations.[25,26] Both of these immunocompromised animals have been previously
established as having the ability to sustain xenograft growth.[25,26] While both SCID-
like pigs have the potential to support tumor growth, the CRISPR/Cas9 derived mutants
are the most logistically sound to utilize given the reliable derivation of the
animals.[25,27,28]
A.4 Materials
A.4.1 Generating RAG2/IL2RG Deficient Pigs
1. Humidified incubator (38.5 °C, 5% CO2, 5% O2).
2. Stereo microscope with warm plate (Nikon).
356
3. Micromanipulator system (Narishige).
4. Microcapillary Puller (Shutter Instrument).
5. FemtoJet (Eppendorf).
6. Glass capillary tubes.
7. Captrol III® micropipet (Drummond Scientific).
8. 100 × 15 mm plastic petri dishes.
9. 35 × 10 mm plastic petri dishes.
10. Manipulation medium: medium 199 supplemented with 0.6 mM NaHCO3, 2.9 mM
Hepes, 30 mM NaCl, 10 ng/ml gentamicin, and 3 mg/ml BSA; pH 7.4.
11. PZM3 culture medium
12. Mineral oil, embryo tested.
A.4.2 Maintaining Cells in Culture
1. Sterile Dulbecco’s Modified Eagle’s Medium (DMEM)
2. Sterile Fetal Bovine Serum (FBS)
3. Sterile Penicillin/streptomycin (optional)
4. Sterile TC vented-cap flasks (25, 75, and 175 cm2 sizes) (see Note 1)
5. Sterile 1x Phosphate Buffered Saline (PBS)
6. 0.25% Trypsin in EDTA
7. 5, 10, 25, and 50 mL serological pipettes
8. 1000 uL pipette tips
9. Incubator (5% CO2)
357
A.4.3 Harvesting Cells for Injection
1. 15 mL conical centrifuge tubes
2. 1000 uL pipette tips
3. 10 uL pipette tips
4. Microcentrifuge tube 1.7 mL
5. Refrigerated centrifuge capable of 125xg
6. Hemocytometer (see Note 2)
7. Trypan blue
8. Matrigel
A.4.4 Sterile Passing in of the Cells
1. Icepack (see Note 3)
2. Small plastic box for spraying in cells
3. Spor-Klenz
A.4.5 Injecting Tumors
1. 1 mL syringes
2. 27-gauge 1/2-inch needles
A.4.6 Monitoring Tumors and Health
1. Plastic Vernier calipers (see Note 4)
2. Rope (8-12 inches long, 1/4 inch diameter) (see Note 5)
3. Health monitoring sheets
358
A.4.7 Ultrasound Imaging (see Note 6)
4. Large container
5. Electronic trimmers for hair removal
6. Laptop Computer
7. SmartUs EXT-1M beamformer (Telemed, Lithunia, EU)
8. AC power cable
9. 100~240 VAC, 50~60 Hz power supply (EN60601-1, Telemed, Lithunia, EU)
10. CTG to USB adaptor cable
11. EchoWave II Imaging software (Telemed, Lithunia, EU)
12. Linear array ultrasound imaging probe (L18-10L30H-4, Telemed, Lithunia, EU)
13. Phased array ultrasound imaging probe (P8-3L10SI-6, Telemed, Lithunia, EU)
14. Sterile water, PBS, or saline
15. Sterile gauze or paper towels
16. Ultrasound jelly
A.4.8 Necropsy
1. Scalpel or sharp, non-serrated knife
2. Gloves
3. Ruler
4. 10% formalin
5. Histology cassettes
6. Cutting board or necropsy table
7. 10% Bleach
359
8. Disposable wypalls or paper towels
A.5 Methods
A.5.1 Generating RAG2/IL2RG Deficient Pigs
1. Conduct in vitro fertilization (IVF) to generate zygotes that will be injected with
CRISPR/Cas9 system targeting RAG2 and IL2RG. (see Note 7)
2. After 2-3 hours post-IVF, wash presumable zygotes in manipulation medium.
3. Place the embryos in a microinjection dish (10-20 µl drop of manipulation
medium covered with mineral oil in 100 x 10 mm petri dish).
4. Inject sgRNAs targeting RAG2 or IL2RG genes and Cas9 mRNA (10 ng/µl
sgRNA and 20 ng/µl Cas9 mRNA are recommended) into cytoplasm of the
presumable zygotes using FemtoJet microinjector under the control of
micromanipulator. Microinjection is conducted on a heating plate at 37 °C. (see
Note 8)
5. After microinjection, wash the embryos twice in PZM3 medium.[29]
6. Place the microinjected embryos in a culture dish (20 µl drops of PZM3 medium
covered with mineral oil in 35 mm dish) and incubate them at 38.5°C, 5% O2, and
5% CO2 in humidified air until the day of embryo transfer.
7. For embryo transfer, identify surrogate sows displaying signs of ovulation. (see
Note 9)
360
8. Transfer the CRISPR/Cas9 injected embryos (about 50-100 embryos depending
on the developmental stage of embryos), cultured for 3-5 days, into the oviduct of
the sows through surgical embryo transfer. (see Note 10)
9. Determine pregnancy by ultrasound at day 30 of gestation.
10. Conduct a gnotobiotic hysterectomy between day 113-115 post fertilization to
derive SCID-like piglets, and then house in gnotobiotic isolators.[30] (see Note
11)
A.5.2 Maintaining Cells in Culture
1. Culture PANC1 cells in DMEM supplemented with 10% fetal bovine serum and
optional 1% penicillin/streptomycin at 37oC and 5% CO2 in tissue culture treated
flasks. (see Note 12)
2. Split cells to keep confluency between 50-80%
a. Aspirate spent media, and wash plate with room temperature PBS.
b. Cover base of the plate with 0.25% trypsin in EDTA, and incubate 5-10
minutes at 37oC and 5% CO2.
c. Add 37oC serum containing culture media, and passage desired
percentage of cells to new flask and add appropriate volume of 37oC fresh
media (i.e. 50ml for a T175).
A.5.3 Harvesting Cells for Injection
361
1. Aspirate culture media, wash with PBS, cover base of the plate with 0.25%
trypsin in EDTA, and incubate 5-10 minutes at 37oC and 5% CO2. Record the
volume of trypsin used.
2. Add serum containing media to deactivate the trypsin and transfer to a 15 ml
conical tube. Record the volume added, which should be 5-8 times the amount of
trypsin used.
3. Transfer 10 uL of cell suspension to a 2mL tube, or smaller, for counting.
4. Centrifuge the 15mL tube at 125xg for 5-10 mins at 4oC.
5. Determine cell concentration using 10ul cells aliquoted with a hemocytometer.
(see Note 13)
6. Remove media from 15mL tube, and resuspend cells in Matrigel for desired cell
concentration and keep in a 1.7 ml tube. (see Notes 14-15)
7. Keep 1.7 mL tube on an ice pack.
8. Aliquot additional 1.7 tube of Matrigel, of equal volume, for control injections.
(see Note 16)
A.5.4 Sterile Passing in of the Cells
1. Place tubes of Matrigel and Matrigel-cells on ice packs in labeled plastic
containers.
2. Set plastic containers inside the gnotobiotic transport isolator port and spray
down with Spor-Klenz.
3. Seal port from transport isolator to two housing isolators with a “Y” cap. Let set
for one hour.[30] (see Note 17)
362
4. Have one individual enter the transport isolator and another enter the home
isolator unit.
5. Open the port openings within both isolators.
6. Carefully, use inside arms to pass plastic container with Matrigel and Matrigel-
cells into home isolator units prior to moving any piglets. (see Note 18)
A.5.5 Injecting Tumors
Figure 0-1: Creating “Tent” to Ensure Subcutaneous Injection. By pulling the pig’s ear forward, a small tent of skin will pull up forming a small tent. To ensure a subcutaneous injection, have the bevel of the needle enter “tent” following a path indicated
1. Pull 0.3-0.6mL Matrigel (without cells) into 1mL syringe. Place 27-gauge 1/2-inch
needle on syringe and return to ice. (see Note 19)
2. Prepare second syringe of Matrigel-cells the same as above, and return to ice.
3. Have handler restrain pig by having the animal rest on one arm, while having
their free hand grasp the pig’s back for stability.
4. Pull the ear of interest away from the site of injection for control Matrigel, creating
a skin-tent (Figure 1). Carefully insert the needle bevel up into this area.
363
5. Inject 100 uL when it is clear the bevel is within the skin-tent, and therefore a
subdermal injection. Pull the needle out in a straight line with a slight wrist-
twisting motion to help avoid any trailing tumor.
6. Once complete, inject the second ear on the same animal before moving on to
the next pig.
7. Make sure that neither injection leaked prior to returning pig to its compartment.
A.5.6 Monitoring Tumors and Health
Figure 0-2: Measuring Subcutaneous Tumor Growth in Gnotobiotic Porcine Isolators. (A) Subcutaneous tumors (indicated by arrow) grown behind the ear can be easily viewed within two days post injection and (B) measured with plastic Vernier calipers. (C) The
1. To measure tumors, have handler restrain the pig by having the animal rest on
one arm, while having their free hand grasp the pig’s back for stability.
2. Gently pull the pig’s ear towards their nose in order to have a clear view of the
injection site, or tumor once growth has been noted.
3. If there is palpable or viewable growth, use calipers to measure the tumor.
Determine the diameter by measuring the widest portion of the growth and the
width perpendicular to that. (Figure 2)
364
4. Calculate diameter by taking the square root of the product of the two numbers
collected above.
5. Once tumor and control injection sites have been monitored, have restrainer hold
the pig by the hind legs.
6. Secure a rope with a slip knot above the pig’s ankle, by sliding the foot through
the opening in the rope and pulling the rope tight.
7. At the other end of the rope, ensure a good grip on the hanging scale.
8. Have the restrainer gently release the pig. With the pig suspended by the
hanging scale, note the weight.
9. Once the weight has been observed and the restrainer regains control of the pig,
remove the rope from the pig’s ankle by loosening the slip knot and sliding the
rope off of the pig’s foot.
A.5.7 Ultrasound Imaging
1. Set up ultrasound imaging system prior to removing pigs from isolators. (see
Note 20)
2. Remove pigs from gnotobiotic isolator by removing both port seals, and having
one person pass pigs out one at a time to another person. (see Notes 21-22)
3. Wash tumor area and shave using electronic trimmers to minimize imaging
interference. (see Note 23)
4. Apply ultrasound jelly to area, and gently press ultrasound probe onto tumor.
5. Use Telemed imaging software to collect desired image.
365
A.5.8 Necropsy
1. Euthanize pigs through desired, IACUC approved methods.
2. Make a posterior cervical incision from the base of the neck to the top of the skull
to allow the skin the be folded outwards.
3. Locate the tumor within the dermis by following the visual markers from the
exterior and palpating the extent of the tumor margins within.
4. Carefully extract the mass. (see Note 24)
5. Make a midline abdominal incision from the base of the ribs down to the pelvic
bone.
6. Carefully remove the vascularized organs that are frequent sites of metastasis:
liver, spleen, and lungs.
7. One at a time preform a bread slice on each of the organs collected to check for
gross metastases.
a. Starting at one end of the organ, make sequential slices each centimeter
until reaching the opposite end of the tissue.
b. Briefly check each tissue for any gross changes in color or shape that
might be indicative of a tumor.
c. Collect any suspect tissue in a histology cassette and store in 10%
formalin.
8. Clean surface with 10% bleach and disposable Wypalls or paper towels when
done.
366
A.6 Notes
1. Our preference is for TC vented-cap flasks; however, utilization of other TC
flasks or plates is based on lab preference.
2. Automatic cell counter can also be used if available.
3. Plastic, sealed ice packs are advised to be used in order to help maintain sterility
when passing cells into gnotobiotic isolators. Whether these are malleable or
solid plastic is based on availability of icepacks and plastic containers for holding
icepacks and cells together.
4. Metal Vernier calipers will corrode after being sterilized for gnotobiotic isolators.
They can be used if necessary, but it is highly advised to utilize plastic as much
as possible.
5. Exact size of rope is not critical. The important part is that the material is smooth
and will not injury the pigs during weighing, and is cut to a length that is long
enough to support two slip knots.
6. Clinical ultrasound imaging systems can be utilized based on availability and
tumor anatomical location. If feasible, using CT or MR’ imaging can also be used
for monitoring orthotopic or metastatic tumor growth.
7. Technical details of the IVF procedures can be found in a previous
publication.[31]
8. The preparation of sgRNA and Cas9 mRNA can be found in a previous
publication.[32]
9. Estrus stage of surrogate sows should be synchronized with developmental
stage of the embryos injected with CRISPR/Cas9 system.
367
10. In vitro-derived pig embryos have low viability. The use of in vivo-derived
embryos may lower the number of embryos transferred to produce SCID-like
pigs.
11. While it is not critical to an oncological study to utilize gnotobiotic housing, for the
health of the SCID-like pigs, it is advised to follow gnotobiotic pig protocols for
hysterectomy delivery and housing to avoid unintending illness.
12. If utilizing an alternative cell line, use appropriate media. For example, if using a
cell line provided by ATCC, use media advised on their website.
13. If it is available, or is a common practice, use an automatic cell counter.
14. We saw successful engraftment with 1.2x106 cells in 100 uL of Matrigel, however
going as high as 6-10x107 cells should also produce viable tumors, but be aware
the increased injection density will increase the initial growth rate.
15. If you use a larger tube, or a larger syringe than advised below (3.4 step 1), it will
be difficult to administer the smaller volume for the injection.
16. Have pairs of tubes designated to go to each gnotobiotic pig isolator.
17. Once the one-hour time has passed, the port is considered sterile. If sterility is
not required (i.e. for a very short study), the incubation time can be shortened.
18. If necessary, pigs can be passed into their home isolators first, and cells passed
in at a later time. However, the pigs will be less effected by anesthesia and more
difficult to inject the further from the hysterectomy that one waits.
19. It is best to not pull too much Matrigel into a syringe at a time, because it will
make it harder to control the injection leading to distorted tumor shapes or miss
calculated injection volumes.
368
20. Monitoring of tumor growth subcutaneously can be done using a linear array
ultrasound (L18-10L30H-4, TELEMED, Lithuania, EU). If you want to image
internal organs for orthotopic tumor placements or to check for metastatic
growths, utilize a phased array ultrasound probe (P8-3L0S1-6, TELEMED,
Lithuania, EU) in order to image a larger, deeper area.
21. If multiple pigs are in the isolator, have a large container to hold the pigs in until
each pig is euthanized individually. The isolator will collapse once it is opened so
you need a container to hold the pigs until they are euthanized.
22. Pass pigs out head first, and they will help by running in the correct direction, into
the arms of the catching handler.
23. To minimize risk of infection to immunocompromised animals when imaging
during study, clean area with sterilized water, PBS, or saline with sterilized
gauze, paper towels, or similar cloth.
24. When extracting the tumor, extracting extra margins can increase the probability
of removing the full tumor if full removal is the priority.
369
A.7 References
1. Aier I, Semwal R, Sharma A, Varadwaj PK (2019) A systematic assessment of
statistics, risk factors, and underlying features involved in pancreatic cancer.
Cancer epidemiology 58:104-110
2. Ghaneh P, Costello E, Neoptolemos JP (2007) Biology and management of
pancreatic cancer. Gut 56 (8):1134-1152
3. Burris Hr, Moore MJ, Andersen J, Green MR, Rothenberg ML, Modiano MR,
Christine Cripps M, Portenoy RK, Storniolo AM, Tarassoff P (1997)
Improvements in survival and clinical benefit with gemcitabine as first-line
therapy for patients with advanced pancreas cancer: a randomized trial. Journal
of clinical oncology 15 (6):2403-2413
4. Davalos RV, Mir L, Rubinsky B (2005) Tissue ablation with irreversible
electroporation. Annals of biomedical engineering 33 (2):223
5. Ivey JW, Latouche EL, Sano MB, Rossmeisl JH, Davalos RV, Verbridge SS (2015)
Targeted cellular ablation based on the morphology of malignant cells. Scientific
reports 5:17157
6. Ivey J, Wasson E, Alinezhadbalalami N, Kanitkar A, Debinski W, Sheng Z, Davalos
R, Verbridge S (2019) Characterization of Ablation Thresholds for 3D-Cultured
Patient-Derived Glioma Stem Cells in Response to High-Frequency Irreversible
Electroporation. Research 2019:8081315
370
7. Martin RC, McFarland K, Ellis S, Velanovich V (2013) Irreversible electroporation in
locally advanced pancreatic cancer: potential improved overall survival. Annals of
surgical oncology 20 (3):443-449
8. Martin RC, Kwon D, Chalikonda S, Sellers M, Kotz E, Scoggins C, McMasters KM,
Watkins K (2015) Treatment of 200 locally advanced (stage III) pancreatic
adenocarcinoma patients with irreversible electroporation: safety and efficacy.
Annals of surgery 262 (3):486-494
9. Bower M, Sherwood L, Li Y, Martin R (2011) Irreversible electroporation of the
pancreas: definitive local therapy without systemic effects. Journal of surgical
oncology 104 (1):22-28
10. Martin II RC, McFarland K, Ellis S, Velanovich V (2012) Irreversible electroporation
therapy in the management of locally advanced pancreatic adenocarcinoma.
Journal of the American College of Surgeons 215 (3):361-369
11. Martin RC (2013) Irreversible electroporation of locally advanced pancreatic head
adenocarcinoma. Journal of Gastrointestinal Surgery 17 (10):1850-1856
12. Saccomandi P, Lapergola A, Longo F, Schena E, Quero G (2018) Thermal ablation
of pancreatic cancer: A systematic literature review of clinical practice and pre-
clinical studies. International Journal of Hyperthermia 35 (1):398-418
13. Hwang JH, Wang Y-N, Warren C, Upton MP, Starr F, Zhou Y, Mitchell SB (2009)
Preclinical in vivo evaluation of an extracorporeal HIFU device for ablation of
pancreatic tumors. Ultrasound in medicine & biology 35 (6):967-975
371
14. Bader KB, Vlaisavljevich E, Maxwell AD (2019) For whom the bubble grows:
physical principles of bubble nucleation and dynamics in histotripsy ultrasound
therapy. Ultrasound in medicine & biology
15. Longo KC, Knott EA, Watson RF, Swietlik JF, Vlaisavljevich E, Smolock AR, Xu Z,
Cho CS, Mao L, Lee FT, Jr., Ziemlewicz TJ (2019) Robotically Assisted Sonic
Therapy (RAST) for Noninvasive Hepatic Ablation in a Porcine Model: Mitigation
of Body Wall Damage with a Modified Pulse Sequence. Cardiovasc Intervent
Radiol 42 (7):1016-1023. doi:10.1007/s00270-019-02215-8
16. Vlaisavljevich E, Owens G, Lundt J, Teofilovic D, Ives K, Duryea A, Bertolina J,
Welling TH, Xu Z (2017) Non-invasive liver ablation using histotripsy: preclinical
safety study in an in vivo porcine model. Ultrasound in medicine & biology 43
(6):1237-1251
17. Vlaisavljevich E, Kim Y, Allen S, Owens G, Pelletier S, Cain C, Ives K, Xu Z (2013)
Image-guided non-invasive ultrasound liver ablation using histotripsy: Feasibility
study in an in vivo porcine model. Ultrasound in medicine & biology 39 (8):1398-
1409
18. Wagner M, Greten FR, Weber CK, Koschnick S, Mattfeldt T, Deppert W, Kern H,
Adler G, Schmid RM (2001) A murine tumor progression model for pancreatic
cancer recapitulating the genetic alterations of the human disease. Genes &
Development 15 (3):286-293
19. Herschkowitz JI, Simin K, Weigman VJ, Mikaelian I, Usary J, Hu Z, Rasmussen KE,
Jones LP, Assefnia S, Chandrasekharan S (2007) Identification of conserved
372
gene expression features between murine mammary carcinoma models and
human breast tumors. Genome biology 8 (5):R76
20. Segatto NV, Remião MH, Schachtschneider KM, Seixas FK, Schook LB, Collares T
(2017) The oncopig cancer model as a complementary tool for phenotypic drug
discovery. Frontiers in pharmacology 8:894
21. Maher B (2013) Tissue engineering: How to build a heart. Nature News 499
(7456):20
22. Flisikowska T, Merkl C, Landmann M, Eser S, Rezaei N, Cui X, Kurome M,
Zakhartchenko V, Kessler B, Wieland H (2012) A porcine model of familial
adenomatous polyposis. Gastroenterology 143 (5):1173-1175. e1177
23. Sieren JC, Meyerholz DK, Wang X-J, Davis BT, Newell JD, Hammond E, Rohret JA,
Rohret FA, Struzynski JT, Goeken JA (2014) Development and translational
imaging of a TP53 porcine tumorigenesis model. The Journal of clinical
investigation 124 (9):4052-4066
24. Schachtschneider KM, Schwind RM, Newson J, Kinachtchouk N, Rizko M,
Mendoza-Elias N, Grippo P, Principe DR, Park A, Overgaard NH, Jungersen G,
Garcia KD, Maker AV, Rund LA, Ozer H, Gaba RC, Schook LB (2017) The
Oncopig Cancer Model: An Innovative Large Animal Translational Oncology
Platform. Front Oncol 7:190. doi:10.3389/fonc.2017.00190
25. Lee K, Kwon D-N, Ezashi T, Choi Y-J, Park C, Ericsson AC, Brown AN, Samuel MS,
Park K-W, Walters EM (2014) Engraftment of human iPS cells and allogeneic
porcine cells into pigs with inactivated RAG2 and accompanying severe
373
combined immunodeficiency. Proceedings of the National Academy of Sciences
111 (20):7260-7265
26. Basel MT, Balivada S, Beck AP, Kerrigan MA, Pyle MM, Dekkers JC, Wyatt CR,
Rowland RR, Anderson DE, Bossmann SH (2012) Human xenografts are not
rejected in a naturally occurring immunodeficient porcine line: a human tumor
model in pigs. BioResearch open access 1 (2):63-68
27. Lei S, Ryu J, Wen K, Twitchell E, Bui T, Ramesh A, Weiss M, Li G, Samuel H,
Clark-Deener S (2016) Increased and prolonged human norovirus infection in
RAG2/IL2RG deficient gnotobiotic pigs with severe combined immunodeficiency.
Scientific reports 6:25222
28. Kang JT, Cho B, Ryu J, Ray C, Lee EJ, Yun YJ, Ahn S, Lee J, Ji DY, Jue N, Clark-
Deener S, Lee K, Park KW (2016) Biallelic modification of IL2RG leads to severe
combined immunodeficiency in pigs. Reprod Biol Endocrinol 14 (1):74.
doi:10.1186/s12958-016-0206-5
29. Yoshioka K, Suzuki C, Tanaka A, Anas IM-K, Iwamura S (2002) Birth of piglets
derived from porcine zygotes cultured in a chemically defined medium. Biology of
reproduction 66 (1):112-119
30. Yuan L, Jobst PM, Weiss M (2017) Gnotobiotic Pigs: From Establishing Facility to
Modeling Human Infectious Diseases. In: Gnotobiotics. Elsevier, pp 349-368
31. Uh K, Lee K (2017) Use of Chemicals to Inhibit DNA Replication, Transcription, and
Protein Synthesis to Study Zygotic Genome Activation. In: Zygotic Genome
Activation. Springer, pp 191-205
374
32. Ryu J, Lee K (2017) CRISPR/Cas9-mediated gene targeting during embryogenesis
in swine. In: Zygotic Genome Activation. Springer, pp 231-244
