ABSTRACT 
 
 
 
 
Title of Dissertation: LIGHT ACTIVATABLE PURE PORPHYRIN 
NANOPARTICLES FOR THE PHOTODYNAMIC 
OPENING OF THE BLOOD-BRAIN BARRIER 
AND GLIOBLASTOMA TREATMENT 
 
  
 Collin Thomas Inglut,  
Doctor of Philosophy, 2022 
  
Dissertation directed by: Assistant Professor, Huang Chiao Huang,  
Fischell Department of Bioengineering 
 
 
 
Glioblastoma (GBM) consistently recurs due to infiltrating cancer cells that cannot be 
removed by surgery and chemotherapy. The diffusive nature of GBM makes complete surgical 
resection unsafe, and the intact blood-brain barrier (BBB) prevents the penetration and 
accumulation of nearly all chemotherapy in infiltrative GBM cells. Existing BBB opening 
strategies are often associated with increased risk of edema, hemorrhage, or neurotoxicity and thus 
have limited clinical success. Photodynamic therapy (PDT) is a photochemistry-based treatment 
modality that has shown promise in treating GBM and opening the BBB in the clinic. In fact, a 
single adjunctive dose of PDT has been shown to add as much as 18 months to patient survival. 
However, the full potential of PDT is limited by the light activation depth of the ?gold standard? 
pro-drug photosensitizer, 5-aminolevulinic acid (5-ALA). In addition, large doses of PDT can 
result in edema and neurotoxicity. To address these issues, our lab has developed a photodynamic 
  
priming (PDP) strategy using the verteporfin (VP) photosensitizer, which operates at low optical 
energy to enhance intratumoral drug accumulation without damaging the healthy brain tissues. 
Unfortunately, VP is hydrophobic and requires liposomal encapsulation for intravenous 
administration, which can alter the photosensitizers cellular pharmaceutics. Here, we develop and 
compare a novel carrier-free pure-photosensitizer nanoparticle to a clinically relevant liposomal 
formulation. 
This dissertation covers a complementary, four-pronged approach to enhance drug delivery 
to brain tumors and treat GBM: (1) Understand the photoactivation depth of clinically relevant 
photosensitizers in the rodent brain for the targeting of infiltrative GBM cells. (2) Explore the 
mechanisms of photochemistry-induced BBB opening. (3) Engineer light-activable 
nanotechnology that can open the BBB, improve drug delivery, and eradicate GBM cells. And (4) 
develop a high-throughput model to examine the BBB integrity and efflux transporter function. 
The central hypothesis of this dissertation is the delivery of photoactivatable pure-photosensitizer 
nanoparticles can eradicate GBM cells and enhance drug delivery to microscopic GBM tumors. 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
 
 
 
 
 
 
 
 
LIGHT ACTIVATABLE PURE PORPHYRIN NANOPARTICLES FOR THE 
PHOTODYNAMIC OPENING OF THE BLOOD-BRAIN BARRIER AND 
GLIOBLASTOMA TREATMENT 
 
 
 
by 
 
 
             Collin Thomas Inglut 
 
 
 
 
 
Dissertation submitted to the Faculty of the Graduate School of the  
University of Maryland, College Park, in partial fulfillment 
of the requirements for the degree of 
Doctor of Philosophy 
2022 
 
 
 
 
 
 
 
 
 
Advisory Committee: 
Assistant Professor, Huang Chiao Huang, Chair 
Associate Professor, Giuliano Scarcelli 
Associate Professor, Kimberly M. Stroka 
Professor, Xiaoming (Shawn) He 
Principal Investigator, Michael M. Gottesman 
Professor, Lawrence C. Washington 
 
 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
? Copyright by 
Collin Thomas Inglut 
2022 
 
 
 
 
 
 
 
 
 
 
 
 
 
  
 
Dedication 
In memory of Jennie Revezzo and Donald J. Inglut, Sr. 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
ii 
 
 
 
Acknowledgements 
There are so many people I would like to thank for making this dissertation 
possible and for supporting me during my time at the University of Maryland. 
 First, I would like to thank my advisor, Dr. Huang Chiao (Joe) Huang. His 
support, drive, and dedication to my learning, has helped me become the best student, 
scientist, and engineer I could be. I am grateful for all his guidance over the past five 
years, and I am thankful to be one of his first Ph.D. students. Thank you, Joe, for truly 
caring about me, both inside and outside of lab.  
 Next, I would like to thank all my lab-mates. First, Barry Liang, who has been 
by my side in lab since day one. Your devotion to science has been an inspiration. I am 
thankful for all the help you have offered me throughout our Huang Lab journey. Going 
on trips to Sushi Para with you were always the best. Thank you, Aaron Sorrin, for your 
team player attitude. Your writing and grammar help is unmatched and truly 
appreciated. Our weekly hangouts during lockdown, watching Tiger King, will be 
missed. Thank you, Brandon Gaitan, for the constant support, help with my research, 
and positive outlook. The fancy steak dinners, cigars on the roof, and horse betting, 
were some of my favorite nights in DC. Our very long days in the dark room will not 
be missed. Thank you all for being amazing lab mates and friends. This work wouldn?t 
have been possible without help from everyone in lab. 
 I would also like to acknowledge several peers and post docs who have helped 
me with collaborative projects. Thank you, Dr. Kelsey Gray, Dr. Nina Connelly, Dr. 
Joanna Thomas, Jill Stabile, and Daniel Najafali. Working and learning with you all 
has been a pleasure.  
iii 
 
 
 
I offer my sincere appreciation to my other advisor and mentor at the NIH, Dr. 
Michael Gottesman and Dr. Rob Robey. Your guidance has helped me become a better 
scientist and critical thinker. I am grateful for the support and your valuable insight into 
my work over the last two years. I deeply enjoyed my time at the NIH with you both. 
Thank you, Fischell Department of Bioengineering staff, for your day-to-day 
help with graduate school requirements. I would especially like to thank Bill Churma, 
for always going out of his way to lend a helping hand and listening to my problems. 
I would like to recognize several friends who have constantly supported me 
during this journey. Thank you to Danial Song, Samm Stewert, Barry Liang, Brandon 
Gaitan, Aaron Sorrin, and Joey Biasi. And a special thank you to my girlfriend, Isabel 
Guterman, who supported me through one of the hardest points in my life. Thank you 
for always listening to me, caring for me, and helping me relax. I am so grateful for 
your endless love, patience, and encouragement.  
Finally, my entire family deserves a huge thank you. My grandparents, Donald 
and Anna, who gave me huge financial support during all my schooling and who always 
believed in me. My hard-working dad and uncle, Don and Tom, who helped install 
their work ethic in me, and who supported and guided me through life. Thank you, 
Tommy, for fixing my car 4 times during grad school when it broke down. And my 
sister, Jessa, who always helped me when times were tough. 
And a final shout out to my cat Papaya, who was always there to sit on my lap 
(and my laptop) while I worked from home. You helped me more than you could know. 
And to my favorite band, Blink-182, who I listened to on repeat, for five years, while 
performing experiments.  The tour is over, I've survived.  
iv 
 
 
 
Table of Contents 
 
 
Dedication ..................................................................................................................... ii 
Acknowledgements ...................................................................................................... iii 
Table of Contents .......................................................................................................... v 
List of Tables ............................................................................................................... ix 
List of Figures ............................................................................................................... x 
List of Abbreviations .................................................................................................... x 
Chapter 1. Introduction ................................................................................................. 1 
1.1 Objective and Significance ................................................................................. 2 
1.2 Approach and Innovation .................................................................................... 3 
1.3 Summary and Dissertation Outline ..................................................................... 5 
Chapter 2. Background ................................................................................................. 7 
2.1 Glioblastoma ....................................................................................................... 7 
2.1.1 Standard of Care .......................................................................................... 7 
2.1.2 The Invasive Front ..................................................................................... 10 
2.1.3 Relevance to Dissertation .......................................................................... 13 
2.2 The Neurovascular Unit .................................................................................... 14 
2.2.1 The Blood-brain Barrier ............................................................................ 16 
2.2.2 Endothelial Cell-Cell Junctions ................................................................. 18 
2.2.3 Endothelial Cell Efflux Transporters ......................................................... 21 
2.2.4 Drug Delivery to the Brain and Clinical Applications .............................. 22 
2.2.5 Relevance to Dissertation .......................................................................... 23 
2.3 Photodynamic Therapy ..................................................................................... 24 
2.3.1 Fundamental Principles ............................................................................. 25 
2.3.2 Photosensitizers ......................................................................................... 28 
2.3.3 Fluorescence Guided Resection ................................................................. 31 
2.3.4 Clinical Applications ................................................................................. 32 
2.3.5 Photodynamic Priming .............................................................................. 33 
2.3.6 Relevance to Dissertation .......................................................................... 35 
Chapter 3. Photodynamic Effect Depth in the Rodent Brain ? ................................... 36 
3.1 Introduction ....................................................................................................... 36 
3.2 Materials and Methods ...................................................................................... 38 
3.2.1 Synthesis and Characterization of Liposomal VP and Liposomal PpIX ... 38 
3.2.2 Photobleaching of Photosensitizers in Ex Vivo Rat Brains ....................... 39 
3.3 Results and Discussion ..................................................................................... 40 
3.3.1 Synthesis and Characterization of Liposomal VP and Liposomal PpIX ... 40 
3.3.2 Evaluation of the Tissue-depth Dependent Photodynamic Effects of VP and 
PpIX .................................................................................................................... 43 
Chapter 4. Understanding the Effects of Photodynamic Priming on Endothelial Cell?
Cell Junctions for Enhancing Drug Delivery ? ........................................................... 49 
4.1 Introduction ....................................................................................................... 49 
4.2 Methods............................................................................................................. 52 
4.2.1 Cell Culture ................................................................................................ 52 
4.2.2 Substrate Coating and Experimental Conditions ...................................... 52 
v 
 
 
 
4.2.3 Cell Viability and Metabolic Activity Assays, and Immunostaining .......... 53 
4.2.4 Immunostaining of Junction Proteins ........................................................ 53 
4.2.5 Microscopy ................................................................................................. 54 
4.2.6 Junction Analysis ....................................................................................... 55 
4.2.7 Western Blot Analysis ................................................................................ 56 
4.2.8 Liposome Synthesis and Characterization ................................................. 57 
4.2.9 Transwell Permeability Assay.................................................................... 58 
4.2.10 Statistical Analysis ................................................................................... 59 
4.3 Results & Discussion ........................................................................................ 60 
4.3.1 Sub-cytotoxic PDP in HBMECs................................................................. 60 
4.3.2 PDP Slightly Alters Cell Morphology and F-actin Organization ............. 63 
4.3.3 PDP Alters the Endothelial Cell-cell Junction Morphology ..................... 66 
4.3.4 PDP Induces HBMEC Permeability to FITC-dextran............................... 71 
4.4 Conclusion ........................................................................................................ 76 
Chapter 5. The Development of Verteporfin Biomolecules for Anti-glioma 
Photodynamic Therapy ? ............................................................................................ 78 
5.1 Introduction ....................................................................................................... 78 
5.2 Methods............................................................................................................. 82 
5.2.1 PSBM Preparation and Purification ......................................................... 82 
5.2.2 Photophysical and Photochemical Characterizations ............................... 83 
5.2.3 Cell Culture ................................................................................................ 84 
5.2.4 Photodynamic Therapy of Glioblastoma Cells Using PSBMs ................... 85 
5.2.5 PSBM Uptake and Localization Studies in Glioblastoma Cells ................ 86 
5.2.6 Immunoblotting .......................................................................................... 87 
5.2.7 Statistical Analyses .................................................................................... 88 
5.3 Results and Discussion ..................................................................................... 88 
5.3.1 Synthesis and Characterization of PSBMs ................................................ 88 
5.3.2 Photoactivity of PSBMs ............................................................................. 91 
5.3.3 Photodestruction of Glioblastoma Cells Using PSBMs............................. 94 
5.3.4 PSBM Uptake ............................................................................................. 96 
5.3.5 Altered Subcellular Localization of PSBM .............................................. 100 
5.4 Conclusion ...................................................................................................... 108 
Chapter 6. Immunological and Toxicological Considerations for Liposome Use ? . 111 
6.1 Introduction ..................................................................................................... 111 
6.2 Toxicological Evaluation of Liposomes and Their Building Blocks .............. 116 
6.2.1 Toxicity of Cationic Liposomes ................................................................ 120 
6.2.2 Interactions with the Mononuclear Phagocyte System ............................ 122 
6.2.3 Effect of Cholesterol Content on the Toxicity of Liposomes .................... 125 
6.3 Activation of the Immune System .................................................................. 127 
6.3.1 Pro-Inflammatory Cytokine Production .................................................. 129 
6.3.3 Hypersensitivity Reactions ....................................................................... 133 
6.3.4 Anti-PEG Response .................................................................................. 135 
6.3.5 Accelerated Blood Clearance .................................................................. 137 
6.4 Conclusion ...................................................................................................... 139 
Chapter 7. Carrier-free, Amorphous Verteporfin Nanoparticles for Enhanced 
Photodynamic Cancer Therapy and Brain Drug Delivery ........................................ 141 
vi 
 
 
 
7.1 Introduction ..................................................................................................... 141 
7.2 Methods........................................................................................................... 143 
7.2.1 NanoVP Synthesis and Characterization ................................................. 143 
7.2.2 Transmission Electron Microscopy ......................................................... 144 
7.2.3 NanoVP Stability, Photoactivity, and Photosensitizer Release Profile ... 144 
7.2.4 NanoVP Electrostatic Stabilization ......................................................... 145 
7.2.5 Cancer Cell Cultures ............................................................................... 146 
7.2.6 Evaluation of Photosensitizer Uptake and PDT Responses In Vitro ....... 146 
7.2.7 Intracellular ROS Production using DCFDA .......................................... 147 
7.2.8 Mitochondrial Membrane Potential using TMRE ................................... 147 
7.2.9 Caspase-3 Western Blots ......................................................................... 148 
7.2.10 Drug Efflux Assay .................................................................................. 148 
7.2.11 In Vivo PDT and Photosensitizer Biodistribution ................................. 149 
7.2.12 Photodynamic Opening of the Blood-brain Barrier in Rodents ............ 150 
7.2.13 Rat Craniotomy ...................................................................................... 151 
7.2.14 Evans Blue Quantification ..................................................................... 151 
7.2.15 Brain Histology ...................................................................................... 152 
7.2.16 Statistical Analyses ................................................................................ 153 
7.3 Results ............................................................................................................. 153 
7.3.1 Formulation and Characterization of NanoVP ....................................... 153 
7.3.2 Photo-activable NanoVP ......................................................................... 158 
7.3.3 In vitro NanoVP Uptake and PDT Efficacy ............................................. 163 
7.3.4 NanoVP Enhances Anti-Glioblastoma PDT ............................................ 169 
7.3.5 NanoVP-PDT Modulation of the Blood-brain Barrier in Rats ................ 172 
7.4 Discussion ....................................................................................................... 174 
7.5 Conclusions ..................................................................................................... 179 
Chapter 8. Development of a High-Throughput Screening Strategy for Discovery of 
Human and Zebrafish ABC Transporter Substrates ................................................. 181 
8.1 Introduction ..................................................................................................... 181 
8.2 Methods........................................................................................................... 184 
8.2.1 Cell Transfection ...................................................................................... 184 
8.2.2 Immunoblotting ........................................................................................ 184 
8.2.3 Cytotoxicity Assays .................................................................................. 185 
8.2.4 High-throughput Substrate Screening ..................................................... 185 
8.3 Results ............................................................................................................. 186 
8.3.1 Establishment and Characterization of HEK293 Cells Stably Expressing 
an ABC Transporter and NanoLuc Luciferase ................................................. 186 
8.3.3 High Throughput Screening of NanoLuc Substrates ............................... 189 
8.4 Discussion and Conclusions ........................................................................... 192 
Chapter 9. Conclusions and Contributions to Science .............................................. 196 
9.1 Verteporfin can be Activated Up to 2 cm in the Rodent Brain ....................... 197 
9.2 PDP-induced Blood-brain Barrier Opening Enhances Drug Delivery ........... 197 
9.3 Combinations of Photosensitizing Biomolecules may increase PDT Efficacy, 
but Limits Cellular Accumulation ........................................................................ 198 
9.4 Pure-drug Carrier-free Nanoparticles of Verteporfin have Superior PDT 
Efficacy Compared to Liposomes ......................................................................... 199 
vii 
 
 
 
9.5 A High Throughput Zebrafish Model aids in the Study of BBB Integrity ..... 200 
9.6 Closing Remarks ............................................................................................. 201 
9.7 Contributions to the Field ............................................................................... 202 
9.7.1 Scientific Contributions ........................................................................... 202 
9.7.2 Peer-Reviewed Journal Publications ....................................................... 204 
9.7.3 National and International Conference Presentations ............................ 206 
Chapter 10. Future Work and Outlook ..................................................................... 207 
10.1 The Treatment of Intracranial Brain Tumors ................................................ 208 
10.2 PDP Synergizes with Irinotecan and Minocycline ....................................... 210 
10.3 A Second-generation Targeted Version of NanoVP ..................................... 213 
10.4 Exploring NanoVP as a Chemotherapeutic .................................................. 214 
10.5 PDP can Modulate ABC Transporter Expression ......................................... 215 
10.6 A Zebrafish Model to Study Blood-brain Barrier Integrity .......................... 218 
10.7 Outlook ......................................................................................................... 219 
Bibliography ............................................................................................................. 221 
 
 
? These chapters were taken or adapted from C.T Inglut?s previously published manuscripts. Specific 
citations and permission from the publishers are included in footnotes at the beginning of each chapter. 
viii 
 
 
 
List of Tables 
 
Table 3.1 Liposomal VP and PpIX ............................................................................. 41 
Table 4.1 Physical Parameters of Rhodamine-liposomes. .......................................... 75 
Table 5.1 PDT Efficiency of Free VP and PSBMs in Glioblastoma Cells. ................ 99 
Table 8.1 GI50 Summary of Transfected HEK293 cells ........................................... 188 
Table 9.1. Summary of Contributions to the field. ................................................... 204 
 
 
ix 
 
 
 
List of Figures 
 
Figure 2.1 Magnetic Resonance Images of Glioblastoma Recurrence ......................... 8 
Figure 2.2 Glioblastoma and the Invasive Front ......................................................... 11 
Figure 2.3 The Neurovascular Unit ............................................................................ 16 
Figure 2.4 Endothelial Cell-Cell Junction .................................................................. 19 
Figure 2.5 Photodynamic Therapy Dosimetry Factors ............................................... 27 
Figure 2.6 ALA mediated Fluorescence Guided Resection of GBM ......................... 32 
Figure 3.1 Absorbance and fluorescence spectra of VP and PpIX ............................. 42 
Figure 3.2 Singlet Oxygen Production and Photobleaching of Liposomal VP and 
Liposomal PpIX .......................................................................................................... 43 
Figure 3.3 Experimental setup for examining the limit of the photodynamic effect .. 45 
Figure 3.4 Depth of Photosensitizer Activation within Brain Tissue Determined via 
Degree of Photobleaching ........................................................................................... 46 
Figure 3.5 Visualization of the Fluorescent Emission of Liposomal VP and Liposomal 
PpIX when Excited at 375 nm .................................................................................... 47 
Figure 4.1 Cell shape factors ...................................................................................... 56 
Figure 4.2 PDP is Sub-cytotoxic to HBMECs ............................................................ 62 
Figure 4.3 Low dose PDP does not Impact Cell Confluency ..................................... 63 
Figure 4.4 PDP Alters Cell Morphology and the Organization of F-actin ................. 65 
Figure 4.5 PDP does not Alter Cell Perimeter ............................................................ 66 
Figure 4.6 PDP Modulates Endothelial Cell-cell Junction Phenotype ....................... 69 
Figure 4.7 Quantitative Analysis of Junction Phenotype. ........................................... 70 
Figure 4.9 PDP does not Alter Total Junction Protein Expression ............................. 71 
Figure 4.8 PDP Impact on Protrusion Length ............................................................. 71 
Figure 4.10 PDP Improves HBMEC Monolayer Permeability to FITC-dextran ....... 73 
Figure 4.11  PDP Improves HBMEC Monolayer Permeability to Rhodamine-
liposomes .................................................................................................................... 76 
Figure 5.1 Synthesis Reaction and Chemical Structure of PSBMs. ........................... 90 
Figure 5.2 MALDI-TOF mass spectrometer analysis of 16:0 Lyso PC-VP and DSPE-
PEG-VP....................................................................................................................... 91 
Figure 5.3 Photophysical and Photochemical Characterization of PSBMs ................ 93 
Figure 5.4 Near-infrared Light Activation of PSBMs Leads to Phototoxicity of 
Glioblastoma cells ....................................................................................................... 95 
Figure 5.5 Cellular Uptake of PSBMs and Free VP ................................................... 98 
Figure 5.6 Comparison of PSBMs and Free VP Uptake in GBM Cell Lines ............. 99 
Figure 5.7 Quantification of VP Fluorescence Signal .............................................. 100 
Figure 5.8 Mitochondrial Localization of Free VP and PSBMs in U251 Cells. ...... 102 
Figure 5.9 Lysosomal Localization of Free VP and PSBMs in U251 Cells ............. 103 
Figure 5.10 Acute Cellular Injury, Lysosomal Membrane Permeabilization, 
Autophagosome Formation, Mitochondrial Membrane Depolarization, and 
Combination Treatment using PSBMs and Free VP ................................................ 106 
Figure 5.11 Acute Cellular Injury, Lysosomal Membrane Permeabilization, 
Autophagosome Formation, Mitochondrial Membrane Depolarization, and 
Combination Treatment using PSBMs and Free VP ................................................ 106 
Figure 6.1 Schematic of a Liposome and its Common Building Blocks .................. 115 
x 
 
 
 
Figure 6.2 Intravenously Injected Liposomes Impact Cancer Progression .............. 125 
Figure 6.3 Schematic Overview of the Complement System Highlighting Two of the 
Main Activation Pathways (Classical vs. Alternative) ............................................. 133 
Figure 6.4 Schematic Overview of the Events that Alter the Pharmacokinetics of 
Repeated Intravenous Doses of Pegylated Liposomes ............................................. 137 
Figure 7.1 Physical Assessment of the Amorphous Nanodrug of Verteporfin 
(NanoVP) .................................................................................................................. 155 
Figure 7.2 Size Quantification of NanoVP TEM Micrograph Images ..................... 156 
Figure 7.3 Representative Ionic Liquid TEM Micrograph ....................................... 156 
Figure 7.4 Alternative NanoVP Synthesis Methods ................................................. 156 
Figure 7.5 Loading Capacity of NanoVP and Liposomal VP .................................. 157 
Figure 7.6 NanoVP Diameter ................................................................................... 157 
Figure 7.7 NanoVP Shelf Life .................................................................................. 158 
Figure 7.8 Probing NanoVP Electrostatic Stabilization ........................................... 158 
Figure 7.9 Photochemical Characterization of NanoVP, Free-form VP, and Liposomal 
VP ............................................................................................................................. 161 
Figure 7.10 High Power Light Activation of VP Results in Limited Photothermal 
Effects ....................................................................................................................... 162 
Figure 7.11 NanoVP Fluorescence Recovers in Physiologically Relevant Mediums
................................................................................................................................... 162 
Figure 7.12 In Vitro PDT Efficacy of NanoVP within U87 Cells ............................ 166 
Figure 7.13 PDT Efficacy within 3T3 Fibroblast Cells ............................................ 167 
Figure 7.14 VP-mediated PDT does not Induce Lysosomal Damage ...................... 168 
Figure 7.15 NanoVP is a Substrate for ABC Transporters ABCG2 and P-gp. ........ 168 
Figure 7.16 NanoVP Dark Toxicity in 3T3 Cells ..................................................... 168 
Figure 7.17 The Phototoxicity and Biodistribution of NanoVP and Liposomal VP in 
an U87 Glioblastoma Xenograft Mouse Model ........................................................ 170 
Figure 7.18 Mouse Weight post PDT Treatment. ..................................................... 171 
Figure 7.19 VP Biodistribution in Mice Organs and U87 Tumors ........................... 171 
Figure 7.20 PDT-induced Blood-brain Barrier Opening within the Rat Brain......... 173 
Figure 7.21 Traditional PDT Damages Healthy Brain Tissue .................................. 174 
Figure 8.1 Immunoblot of HEK293 Transfected Cell Lines .................................... 187 
Figure 8.2 Cytotoxicity Screening within Transfected HEK 293 cells .................... 188 
Figure 8.3 Bioluminescence Signal of Native Coelenterazine and its Analogs ....... 189 
Figure 8.4 ABC Substrate Screening ........................................................................ 191 
Figure 8.5 Bioluminescence Signal Percent Increase ............................................... 192 
Figure 10.1 NanoVP-mediated Interstitial PDP Treatment does not Result in Healthy 
Tissue Damage .......................................................................................................... 209 
Figure 10.2 Triple Combination Treatment within U87 Cancer Cells ..................... 211 
Figure 10.3 In Vivo Treatment Timeline .................................................................. 213 
Figure 10.4 GBM39 Normalized Tumor Volume .................................................... 213 
Figure 10.5 NanoVP-mediated PDP and YAP/TAZ Inhibition for GBM Killing ... 215 
Figure 10.6 PDT Dose Viability Curves ................................................................... 217 
Figure 10.7 Immunoblots of ABCG2 within Transfected HEK293 cells ................. 218 
Figure 10.8 Native coelenterazine and coelenterazine-h are transported by Abcb4 or 
Abcg2a at the zebrafish BBB .................................................................................... 219 
xi 
 
 
 
 
List of Abbreviations 
 
16:0 Lyso-PC ? 16:0 lysophosphocholine 
ABC ? ATP-binding cassette 
ALA ? aminolevulinic acid 
ABCB1- ATP Binding Cassette Subfamily B Member 1 
ABCG2 ? ATP binding cassette subfamily G member 2 
BBB ? blood-brain barrier 
CPT-cAMP ? 8-(4-chlorophenylthio) adenosine-3?,5?-cyclic monophosphate sodium 
DOPE ? dioleoylphosphatidylethanolamine  
DOTAP ? N-(2,3-Dioleoyloxy-1-propyl)trimethylammonium methyl sulfate  
DSPE-PEG ? distearoylphosphoethanolamine-polyethylene-glycol 
EGFR ? epidermal growth factor receptor  
FCCP ? carbonilcyanide p-triflouromethoxyphenylhydrazone  
FDA ? United States Food and Drug Administration 
FGR ? fluorescence-guided resection  
FITC-dextran ? fluorescein isothiocyanate?dextran  
GBM ? glioblastoma  
H&E ? haematoxylin and eosin  
HBMECs ? human brain microvascular endothelial cells 
HSR ? hypersensitivity reactions  
IC50 ? half-maximal inhibition concentration  
IL ? interleukin  
JAnaP ? Junction Analyzer Program  
LD50 ? median lethal dose  
MGMT ? O6-methylguanine methyl transferase  
MMP ? metalloproteinase proteins 
MPS ? mononuclear phagocyte system 
MTT ? (4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide  
PBS ? phosphate buffered saline 
xii 
 
 
 
PC ? phosphatidylcholine 
PDP ? Photodynamic priming 
PDT ? photodynamic therapy 
PE ? phosphatidylethanolamine 
PEG ? polyethylene glycol 
PpIX ? protoporphyrin IX 
PS ? phosphatidylserine 
RIPA ? radioimmunoprecipitation assay 
ROS ? reactive oxygen species 
SOSG ? singlet oxygen sensor green 
SEM ? standard error of the mean 
TAMs ? tumor-associated macrophages 
TEM ? transmission electron microscopy 
TMRE ? tetramethylrhodamine ethyl ester 
TMZ ? temozolomide 
VE-cadherin ? vascular endothelial cadherin 
VP ? verteporfin 
ZO-1 ? zonula occluden-1  
xiii 
 
 
 
1. Chapter 1. Introduction 
 
Glioblastoma (GBM) is the most common and deadly brain cancer. It is 
characterized by highly infiltrative cells that invade away from the primary tumor and 
into the healthy brain. Patients with GBM have a near 100% recurrence rate, with 80% 
of the tumors recurring within 1-2 cm of the original tumor edge [1, 2].  GBM is 
typically treated with surgical resection followed by chemotherapy and radiation to 
eradicate the residual invasive cells [3]. However, complete surgical resection of GBM 
is near impossible and unsafe. In addition, most patients will experience resistance to 
chemotherapy or discontinue maintenance treatment due to complications [4, 5]. While 
the systemic toxicity of chemotherapy can be reduced by nanoparticle systems [6], they 
do not effectively transverse the intact blood-brain barrier (BBB), diminishing their 
treatment efficacy [7].  
Several strategies have been developed to disrupt the continuous tight junction 
network that joins the adjacent endothelial cells for enhanced drug delivery, including 
the infusion of hyperosmolar solutions and focused ultrasound [8]. However, these 
methods are not yet universally available in the clinic due to a number of limitations, 
such as lack of selectivity or healthy tissue damage [8-11]. Moreover, many of these 
strategies focus on endothelial tight junction disassociation and do not account for BBB 
closing nor their cytotoxicity on the other supportive cells, including astrocytes and 
neurons [8]. 
Photodynamic therapy (PDT) has shown promise in opening the BBB and 
treating the deadliest brain cancer in adults- GBM. In fact, a single adjunctive dose of 
PDT has been shown to add up to 18 additional months to patients? lives [12]. 
1 
 
 
Traditionally, PDT involves the high optical energy (80-300 J/cm2) activation of non-
cytotoxic photosensitizers to generate reactive oxygen species that can kill cancer cells 
[13]. PDT requires careful optimization as high doses within the brain can induce 
edema and neurotoxicity [14-20]. Photodynamic priming (PDP), a subtherapeutic dose 
of PDT, using the verteporfin (VP) photosensitizer can modulate the vasculature and 
enhance intratumoral chemotherapy accumulation without damaging the surrounding 
healthy tissues [19]. However, VP is insoluble in water and requires a drug delivery 
vehicle. The current gold standard for delivering VP to patients is Visudyne, a Food 
and Drug Administration (FDA) approved liposome [21]. Still, liposomes can limit 
cancer cell uptake, which may diminish treatment efficacy. It is also well recognized 
there are many immunological and toxicological considerations for the design and use 
of liposomal drugs [6]. A pure-drug carrier-free nanoparticle of VP that resembles the 
free-form version of VP could be ideal for PDP and GBM treatment.  
1.1 Objective and Significance 
The overall objective of this dissertation is to examine how photochemistry 
can be leveraged to enhance drug delivery to the brain and ultimately improve the 
treatment efficacy for the deadliest brain cancer, GBM. We accomplished this goal 
by engineering a novel drug delivery system and repurposing a traditional 
photochemistry-based treatment modality, PDT. We then applied these tools to 
understand the mechanisms behind inducing GBM cell death and BBB modulation. 
The significance of this dissertation lies in: (1) A unique photodynamic priming 
strategy, fundamentally different from the conventional GBM treatment, to selectively 
open the BBB for enhanced delivery of therapeutics; (2) The development of a pure-
2 
 
 
drug carrier-free nanoparticle of the hydrophobic photosensitizer VP; (3) It focuses on 
the underlying cause of recurrence for the most common and lethal brain tumor, the 
infiltrative GBM cells beyond the margins of surgical resection. 
1.2 Approach and Innovation 
This dissertation focuses on engineering a drug delivery system and designing 
a photochemistry-based treatment strategy that can be readily inserted into the existing 
clinical workflow for GBM. We will first examine the photosensitization reaction 
efficiency and activation depth of two clinically relevant photosensitizers in an ex vivo 
rodent brain model. We will investigate how to better deliver the hydrophobic 
photosensitizer VP to patients by engineering and thoroughly characterizing multiple 
nanoparticle systems, including biomolecule-VP conjugates, Liposomal VP, and a pure-
drug carrier-free nanoparticle of VP. We seek to understand how the different 
nanoparticle systems induce GBM cell death and determine their treatment efficacy 
compared to the free-form photosensitizer, which cannot be used clinically. We will 
further explore how the sub-cytotoxic PDT can open the BBB for the delivery of 
therapeutics to the infiltrative GBM cells beyond the surgical margins and examine the 
impact on healthy brain tissues. The central innovative elements of this dissertation 
are:  
1) Sub-cytotoxic Photodynamic Priming to Modulate the Blood-brain Barrier 
The concept of using the VP photosensitizer, which has a longer excitation 
wavelength (690 nm), over the clinical ?gold standard? and ?tumor cell-targeted? 
prodrug aminolevulinic acid (ALA)-induced protoporphyrin IX (PpIX; 635 nm), to 
open the BBB is new [22]. The deeper penetrating light/ photosensitizer activation and 
3 
 
 
the ability to target the vasculature (on-demand and based on the selected drug-light 
interval) will allow for a more selective and effective opening. The spatiotemporal 
nature of PDT, confining light to the specified treatment areas, can also reduce normal 
tissue damage [23]. Mechanistically, we will examine how sub-cytotoxic PDT, known 
as priming, modulates the cell-cell junctions within brain endothelial cells for enhanced 
drug delivery. We will investigate the efficacy of VP- and ALA-mediated BBB opening 
in rodents. 
2) A Pure-drug, Carrier-free Formulation of the verteporfin Photosensitizer 
Liposomes hold great potential as drug delivery vehicles due to their 
biocompatibility and modular properties [24, 25], coupled with the significant 
advantage of attenuating the risk of systemic toxicity from the encapsulated 
chemotherapy agent [26-28]. However, there are still several drawbacks that limit their 
range of applications. Some of the biggest drawbacks of liposomes are the ?Accelerated 
Blood Clearance Phenomenon?, the redirection of the therapeutic payload to 
mononuclear phagocyte system containing organs (i.e., liver and spleen), and the 
diminished cellular uptake [29]. Repeated injections of pegylated liposomes can lose 
their ?stealth? ability and are more quickly cleared from the blood due to the abundant 
secretion of anti-PEG IgM, IgE, and immunoglobulin G, followed by the opsonization 
of C protein fragments (complement activation), and finally uptake by macrophages 
[30-43]. To minimize the patient?s exposure to liposomes, we engineered a pure-drug, 
carrier-free formulation of VP, termed NanoVP. Analysis revealed NanoVP had 
superior in vitro and in vivo cancer treatment efficacy compared to a liposomal 
formulation that represents the current gold standard. We further examined the 
4 
 
 
mechanisms of cancer cell death for different drug delivery systems of VP, including 
our novel NanoVP system. 
1.3 Summary and Dissertation Outline 
This dissertation is organized as follows: Chapter 2 provides an overview of 
GBM, the BBB, and PDT. This chapter focuses on current treatments for GBM while 
highlighting our understanding of why the disease is so deadly; The infiltration GBM 
cells that invade outwards into the healthy brain and the defense mechanisms of the 
BBB that limit drug penetration into the brain. An overview of PDT is provided with a 
focus on how it is currently applied in the clinic for GBM treatment and the barriers 
that need to be overcome for it to be more effective. Chapter 3 explores the activation 
depth of two clinically relevant photosensitizers in the rodent brain. Here, singlet 
oxygen production capabilities are correlated with photosensitizer photobleaching to 
determine the ideal photosensitizer for PDT treatment of GBM. Chapter 4 examines 
the mechanisms governing photochemistry-induced BBB opening. Non-optimized and 
high doses of PDT within the brain can result in healthy tissue damage. In this chapter, 
we examine the impact of sublethal PDT doses on human brain microvasculature 
endothelial cells and cell-cell junction maturity. We further examine the ability to 
deliver nanoparticles past the endothelial cells in a transwell model. Chapter 5 focuses 
on developing VP-biomolecule conjugates for PDT treatment. The three VP-
biomolecule conjugates are thoroughly characterized in situ and in vitro to determine 
the impact of covalent conjugation on VPs photochemical properties and PDT efficacy. 
Chapter 6 investigates the immunological and toxicological considerations for the 
design of liposomes. While the advantages of liposomes are well understood, this 
5 
 
 
chapter explains the several drawbacks that limit their range of clinical applications. 
We aim to shed light on key considerations that should be factored into the design and 
clinical use of liposomal formulations. Chapter 7 spotlights the development of a pure-
drug carrier-free nanoparticle of VP. This chapter covers synthesis methods and 
examines the novel nanoparticle's photochemical properties in storage and 
physiologically relevant environments. This chapter further explores the biomedical 
applications of the pure-drug nanoparticle in vitro and in vivo. PDT efficacy is 
examined within a subcutaneous GBM mouse model and the degree of photochemistry-
induced BBB opening is determined using an intracranial rat model. Two clinically 
relevant photosensitizer formulations are used for comparison. Chapter 8 focuses on 
developing a transgenic zebrafish model for studying BBB integrity and drug efflux 
transporter function. A high throughput cell-based assay is developed to determine if 
luciferase substrates are also substrates for the zebrafish drug transporters present at 
the BBB. 
 
 
 
 
 
 
 
 
 
6 
 
 
2. Chapter 2. Background 
2.1 Glioblastoma 
GBM is the most common and deadly brain tumor, accounting for more than 
77% of all primary malignant tumors of the central nervous system. There are 
approximately 12,000 new cases annually in the US [1, 2]. The survival rates remain 
almost unchanged in decades, with the median survival being 14-16 months and the 5-
year survival rate under 5%. Patient survival without treatment is only 6 months [1, 2]. 
Even with the world?s best healthcare and medical treatments at his disposal, the late 
Senator John McCain succumbed to the disease in August 2018 after battling it for 13 
months. 
2.1.1 Standard of Care 
GBM recurrence is nearly universal due to the inability of the current treatment 
to remove and eradicate the entire cancerous cell population (Figure 2.1) [44, 45]. 
Current first-line treatment includes maximal surgical resection followed by 
chemotherapy (temozolomide, TMZ, 75 mg/m2 daily) and radiation (~1.8?Gy daily, 
~30 fractions total) for 6 weeks, followed by 6 months of adjuvant chemotherapy 
(TMZ, 150 mg/m2 for 5 days every 28 days) [3]. Surgery is performed to remove the 
bulk of the primary tumor; however, because the tumor preferentially invades the 
healthy brain tissue along myelinated fibers in white matter tracts and blood vessels, 
complete surgical removal is unsafe [46-49]. In fact, studies have shown primary tumor 
size is not correlated with survival outcomes [50], but rather if the location of the tumor 
is near a major white matter tract (e.g., the cortico-spinal tract and the anterior thalamic 
radiation) [51].  
7 
 
 
Figure 2.1 Magnetic Resonance Images of Glioblastoma Recurrence. (a) Fluid-attenuated 
inversion recovery image prior to treatment. (b) Post-contrast T1-weighted image prior to 
treatment. (c) Post-contrast T1-weighted image post-surgery. (d) post-contrast T1-weighted 
image after the completion of radiotherapy. (e) Post-contrast T1-weighted image at 10 months 
after diagnosis. (f) Post-contrast T1-weighted image at recurrence was fused with the planning 
computed tomography image, revealing local recurrence. The blue line indicates the 54-Gy 
IDL, the green line indicates the 60-Gy IDL and the red line marks the tumor recurrence 
volume. IDL, isodose line. Reprinted from Zhou et al., Recurrence patterns in patients with 
high-grade glioma following temozolomide-based chemoradiotherapy. Mol Clin Oncol 5(2): 
289-294. 2016. Permission obtained from Spandidos-publications. 
 
The chemotherapy TMZ is used to eradicate the remaining infiltrative 
cancerous cells after surgery. TMZ is a DNA alkylating agent that induces cell cycle 
arrest at G2/M by delivering a methyl group to purine bases of DNA [52]. However, 
approximately 50-55% of newly treated patients will have insignificant or no response, 
known as intrinsic drug resistance, and about 40% will later develop resistance to the 
8 
 
 
therapy, known as acquired drug resistance [4, 5]. The DNA repair protein O6-
methylguanine methyl transferase (MGMT) removes the added methyl groups from the 
O6 position of guanine in DNA, stopping eventual DNA strand breaks [53]. The 
upregulation of this resistance mechanism is correlated with a worse patient prognosis 
and lower median patient survival [54-56]. For example, patients with methylated 
MGMT promotors (i.e., decrease in MGMT protein) had a 2-year survival rate of 49%, 
while patients with unmethylated MGMT promotors had a 2-year survival rate of 15% 
[54].  
Large doses of TMZ are required to reach therapeutic levels in the brain and 
treatment GBM as only 17.8 ? 13.3% can cross the intact BBB [57]. One method to 
combat drug resistance and the high expression of MGMT is to increase the dose of 
TMZ. When MGMT repairs the alkylated DNA, it becomes irreversibly inactivated 
when the alkyl group is transferred to the cysteine moiety in its active site. Dose 
escalation has the potential to increase the rate of DNA alkylation and outpace the rate 
of MGMT protein synthesis [53]. However, dose-escalation studies have shown to 
induce hematologic toxicity. In one study that examined an alternative dosing schedule 
with an increased amount of TMZ (regimen:7-days-on/7-days-off,150 mg/m2/day), 
~38% of patients required a dose reduction due to thrombocytopenia and/or 
neutropenia [58]. Moreover, hematologic toxicity is not uncommon with conventional 
TMZ dosing. A retrospective review (conducted at Sidney Kimmel Comprehensive 
Cancer Center at Johns Hopkins, Baltimore, MD) found grade 3?4 thrombocytopenia 
occurred in 19% of patients and grade 3?4 neutropenia or anemia occurred in at least 
8% of patients. Of the 10 patients that experienced grade 3?4 thrombocytopenia, 9 
9 
 
 
patients (17% of total) discontinued treatment [59]. A pooled analysis from 4 
randomized trials for newly diagnosed GBM showed only 291 of 624 (46.6%) patients 
continued with maintenance TMZ (150 mg/m2 for 5 days every 28 days) [60].  Lastly, 
TMZ is known to cause healthy brain tissue damage. Although hard to study in humans, 
rats treated with TMZ (25 mg/kg) experienced a 34% decrease in new cells generated 
in the granule cell layer within the hippocampus [61]. While TMZ has numerous 
drawbacks, including treatment efficacy, hematologic toxicity, and healthy brain 
damage, there is a reliance on it for first-line treatment due to its ability to accumulate 
within the brain [62]. Other chemotherapeutics and treatment options are limited 
because of the BBB, which restricts drug penetration into the brain.  
 When the tumor recurs or when the patient stops responding to the initial 
treatment, different salvage therapies are used, although there is no standardized 
regimen. Due to the diffuse nature of the tumor, only 25% of patients are eligible for a 
second surgery [63]. If another surgery is performed, biodegradable chemotherapy 
wafers containing carmustine can be placed in the resection cavity [64]. Other options 
include the vascular endothelial growth factor receptor inhibitor cediranib, the anti-
vascular endothelial growth factor monoclonal antibody, bevacizumab [65], re-
irradiation [66], and other chemotherapeutics (e.g., irinotecan and Liposomal 
irinotecan) [67].  
2.1.2 The Invasive Front 
As evident by the local recurrence patterns, the infiltrating cancer cells that 
invade the healthy brain tissue are usually located within 1-2 cm from the original 
lesion and result in an ~80% local recurrence rate (Figure 2.2) [44, 45]. Less than 2% 
10 
 
 
of patients with GBM have metastasis beyond the brain, which tends to occur in late 
stage diseases and after surgery [68]. The infiltrative cells that invade the healthy brain 
are difficult to treat due to the inability to safely remove them all during surgical 
resection and the inability to effectively deliver chemotherapy to them because of the 
intact BBB [69]. Genomic analyses have revealed invasive glioma cells and recurrent 
tumors are distinctly different from the primary tumor, usually as a result of treatment. 
For example, the primary tumor core is known to contain glioma stem cells enriched 
with CD133+ that resemble proneural cells. In contrast, glioma cells on the invasive 
edge are enriched for CD109+ and resemble a mesenchymal subtype [70]. 
 
Figure 2.2 Glioblastoma and the Invasive Front. (a) H&E-stained cortical section of 
glioblastoma. (b) IDH1 R132H immunostaining highlights the neoplastic cells. (c,d) IDH1 
R132H-positive neoplastic cells at the tumor edge and centimeters away into the healthy brain. 
Reprinted from Olar and Aldape. Using the molecular classification of glioblastoma to inform 
personalized treatment. J. Pathol., 232: 165-177, 2013. Permission obtained from Elsevier and 
Copyright Clearance Center?s RightsLink?. 
11 
 
 
There are numerous molecular mechanisms and positive feedback loops 
underlying GBM motility. Some of the well-understood factors include tumor stiffness 
and hypoxia. Tumor stiffness, which is upregulated by the production of collagen, 
fibronectin, hyaluronic acid, and laminin, promotes glioma cell migration and is 
correlated with glioma progression [71]. Studies have shown that GBM cell lines 
spread extensively and migrate rapidly on highly rigid extracellular matrixes. Less rigid 
extracellular matrix comparable to typical brain tissue result in rounded and stationary 
cells [72]. The glioma?s stiff extracellular matrix promotes invasion by upregulating 
promigratory factors. For example, the overexpression of hyaluronic acid in the tumor 
microenvironment promotes cell invasion through positive feedback regulation of 
nuclear factor-?B, a transcription factor that regulates cancer cell proliferation, 
differentiation, motility and survival [73]. The activation of nuclear factor-?B is linked 
to stem-like cell maintenance, epithelial-to-mesenchymal transition, promotion of 
angiogenesis, and resistance to radiotherapy [74]. Additionally, in response to a stiff 
extracellular matrix, glioma cells will upregulate matrix metalloproteinase proteins 
(MMP), secreted or membrane-anchored endoproteinases, to degrade and remodel their 
environment. By destroying their surrounding environment, glioma cells can invade 
and migrate. Commonly, MMP-9 is not detected in healthy brain samples, but active 
MMP-9 is found in 50% of tumors [75]. MMPs create positive feedback loops with 
other pro-migratory factors. MMP-2 and MMP-9 will active the dormant pro-migratory 
transforming growth factor-?, which stimulates the production of more MMP-2 [76]. 
Transforming growth factor-? is known to promote an epithelial-to-mesenchymal 
transition in cancerous cells. After receiving signals or sensing factors including, 
12 
 
 
transforming growth factor -? or hypoxia from other cancerous cells, glioma cells will 
begin to migrate or invade outwards from the primary tumor [71]. They will transition 
from an epithelial to mesenchymal state, detach from the primary tumor by recycling 
(i.e., endocytosing) their integrin and cadherin connections, remodel their microtubule 
network (i.e., form a leading edge), and upregulate new integrins that form links with 
the existing surrounding structures such as ?9?1 and CD44 [77].  
Low tumor oxygenation or hypoxia results from microvascular hyperplasia, an 
exacerbated form of angiogenesis. The secretion of proangiogenic factors, such as 
vascular endothelial growth factors and interleukin-8 (IL-8), results in the hyper-
proliferation and recruitment of endothelial cells and, ultimately, the rapid growth of 
defective and permeable blood vessels. The collapse of these blood vessels yields 
hypoxic foci throughout the tumor. The diminished nutrients promote glioma migration 
and invasion [78]. Additionally, the overexpression of the proangiogenic factors, is 
correlated with poor prognosis in glioma patients [79]. 
2.1.3 Relevance to Dissertation 
The focus of this dissertation is on the treatment of brain cancer. The glioma 
cells that invade the healthy brain are the main source of disease recurrence, which 
elude current treatment options [44, 45]. As such, understanding why current treatment 
options are not effective against the invasive cells and the mechanisms behind glioma 
invasion will help govern therapeutic discovery and implication. Surgical resection of 
all the invasive cells is not feasible and unsafe [80]. Surgical recession is stopped as 
soon as functional brain tissue is reached [80]. On the other hand, the intact BBB 
diminishes the efficacy of the follow-up chemotherapy regimen [7]. This 
13 
 
 
simultaneously drives the need to develop a treatment strategy that can directly kill the 
glioma cells during surgery and synergize with the chemotherapy regimen.  
In this dissertation, the adjunctive treatment modality, PDT, and the new drug 
delivery system, pure-drug carrier-free nanoparticles of VP, have been the focus of 
study for treating GBM. Another technique to treat GBM relies on another 
photosensitizer, pro-drug ALA, to aid in the surgical removal of the primary tumor. 
However, this surgical technique does not focus on the invasive cells. The following 
sections will discuss this surgical technique in detail and examine the limitations we 
seek to overcome. While our focus here is on treating GBM, we believe the developed 
drug delivery system is relevant to other therapeutics and cancers. Chapter 7 will focus 
on developing our nanoparticle and discussing its other biomedical applications. 
2.2 The Neurovascular Unit 
The brain is one of the most vascularized organs in the body. The neurovascular 
unit, composed of endothelial cells, neural cells (i.e., astrocytes, neurons), pericytes, 
microglia, and the basement membrane, regulates nutrient transport, maintains brain 
homeostasis and protects the central nervous system [81-83]. Nutrients are carried up 
to the brain through large cerebral arteries, which consist of smooth muscle cells and 
endothelial cells, with neurons and astrocytes nearby. The cerebral arteries branch into 
smaller pial arterioles, which consist of endothelial cells, smooth muscle cells, 
meningeal cells, and perivascular nerves that run along the surface of the brain. Pial 
arterioles then branch into penetrating arterioles, which penetrate through the fluid-
filled Virchow-Robin space and into the brain. The intracerebral arterioles, which 
contain a single layer of smooth muscle cells, will then branch into brain capillaries, 
14 
 
 
known as the BBB (Figure 2.3) [84]. Besides maintaining a high degree of 
permeability, the neurovascular unit regulates blood flow, angiogenesis, neurogenesis, 
and neurotransmitter turnover [85]. For example, glutamate, an excitatory 
neurotransmitter, is released by neurons within the brain. However, large 
concentrations of glutamate can be damaging to nerve cells. Glutamate is normally 
taken up by astrocytes through the EAAT1 transporter. Glutamate is either metabolized 
to ?-ketoglutarate by glutamate-dehydrogenase and used within the tricarboxylic acid 
cycle or converted into glutamine by glutamine synthetase and shuttled back over to 
neurons [84]. However, damage to cells within the neurovascular unit can disrupt this 
cycle. Instead of astrocyte taking up glutamate, they can secrete it, leading to neuron 
cell death [86]. When designing methods to open the BBB, it is critical to avoid severe 
damage to cells of the neurovascular unit. 
15 
 
 
Figure 2.3 The Neurovascular Unit. (Left) Diagrams of the intraparenchymal arterioles and 
the capillaries within the brain.  (Right) Lipophilic dye DiO was used to visualize the 
vasculature (green). CD206 immunocytochemistry was used to visualize the macrophages 
(blue) located in the perivascular and subarachnoid space. Reprinted from Iadecola, The 
Neurovascular Unit Coming of Age: A Journey through Neurovascular Coupling in Health and 
Disease. Neuron 96(1): 17-42., 2017. Permission obtained from Elsevier and Copyright 
Clearance Center?s RightsLink?. 
 
2.2.1 The Blood-brain Barrier 
The BBB is a network of continuous non-fenestrated capillaries within the 
neurovascular unit. There are two main cell types that form the BBB: a single layer of 
microvascular endothelial cells that forms the vessel well and supporting mural cells 
that include pericytes and astrocytic foot processes [83]. The endothelial cells and 
pericytes secrete a vascular basement membrane (i.e., extracellular matrix) comprised 
of type IV collagen, laminin, and other glycoproteins, that aids in signaling processes. 
16 
 
 
The BBB forms a physical and chemical barrier between the circulatory and central 
nervous systems, tightly regulating the influx and efflux of biological substances for 
the brain parenchyma [7, 83]. Nutrients required for proper brain function enter the 
brain via transporter proteins, carrier-specific transcytosis, diffusion across the 
endothelial cells, and extremely restricted paracellular diffusion [87].   
Compared to other endothelial cells throughout the body, the endothelial cells 
of the BBB are characterized by the presence of inter-endothelial tight junctions and 
the lack of caveolae at the luminal surface [88]. The cell-to-cell junctions that join the 
adjacent endothelial cells greatly limit the paracellular flux of solutes, forcing most 
molecules to undergo receptor-mediated endocytosis and transcytoses. For example, 
the BBB restricts and regulates the movement of ions and fluid, specifically K+ and 
Ca2+, through transporters, to ensure the brain interstitial fluid is optimal for neuronal 
function [89]. 
Pericytes sit on the abluminal surface of the endothelial tube and are identified 
by Platelet-derived growth factor receptor beta (PDGFR-?) and neuron-glia antigen 2 
(NG2) expression [90]. Their proximity to the endothelial cells allows for ion, 
metabolite, and messenger exchange. Their phagocyting capabilities aid with the 
removal of toxic metabolites. Similar to smooth muscle cells, pericytes possess 
contractile characteristics. In response to depolarization or neurotransmitter action, 
their contractile proteins allow them to control the diameter of the capillary and regulate 
blood flow [88]. 
Astrocytes are the most abundant cell type in the CNS. Their endfeet encases 
the vascular tube, forming an interface between the vascular and the neuroglia in the 
17 
 
 
neurovascular unit. This neurovascular union allows astrocytes to relay signals to 
pericytes that regulate blood flow. Their main function is to help maintain the brain's 
ionic, amino acid, neurotransmitter, and water concentrations [91]. Numerous studies 
have shown the importance of astrocytes in BBB formulation as well as the ability of 
purified astrocytes to provide similar BBB features to other blood vessel types [92].  
2.2.2 Endothelial Cell-Cell Junctions 
Adherens, tight, and gap junction proteins govern endothelial cell-to-cell 
interactions. The over 40 proteins within the cell-cell junction are responsible for cell 
adhesion, barrier formation, and the exchange of nutrients between cells (Figure 2.4) 
[93]. Transmembrane proteins create junction complexes with identical proteins on 
adjacent cells at sites of cell-to-cell contact and are tethered to the actin cytoskeleton 
by various intracellular components (Figure 2.4) [94]. Proper formation of the junction 
complexes is crucial for vascular function as cell-cell interactions regulates 
angiogenesis, cell migration, and cell polarity [94]. Diminished junction integrity has 
been associated with numerous diseases including, cancer, stroke, ischemia, 
atherosclerosis, and many neurodegenerative diseases [95]. 
18 
 
 
Figure 2.4 Endothelial Cell-Cell Junction. (top panel) Endothelial cells create a barrier 
between the circulatory and central nervous systems, which are exposed to constant fluid shear 
stress. (bottom panel) Transmembrane proteins that form the cell-cell junction. Tight and 
adheren junction proteins are responsible for cell adhesion and barrier formation. The 
transmembrane proteins are connected to the actin cytoskeleton via scaffolding proteins. 
Reprinted from Cerutti and Ridley, Endothelial cell-cell adhesion and signaling. Experimental 
Cell Research 358(1): 31-38. 2017. Open access. Permission given by Elsevier (Creative 
Commons CC-BY license). 
 
 
Adherens junctions initiate and maintain cell-to-cell contact, a critical first step 
in tight junction formation. Adherens junctions are primarily formed by cadherin 
proteins, with vascular endothelial (VE)-cadherin being strictly expressed in 
endothelial cells. Cadherin adhesion relies on homophilic, Ca2+-dependent interactions 
at the extracellular domains [96]. The intracellular domains, which contain binding 
19 
 
 
locations for transcription factors, are connected to the cytoskeleton via ?- catenin or 
?-catenin. The transcription factor p120 also binds to this domain, which prevents 
endocytosis of the cadherins via inhibited translocation to the nucleus [97]. 
Translocation of p120 to the nucleus initiates clathrin-dependent endocytosis and 
increases barrier permeability. Another pathway to modulate cadherin-mediated 
permeability is through increased cellular contraction. Phosphorylation of myosin light 
chain results in the activation of p21-activated kinase and increased contraction [97]. 
Lastly, ve-cadherin can diminish the angiogenetic effects of vascular endothelial 
growth factor by inhibiting the internalization and signaling of vascular endothelial 
growth factor receptor-2 [98]. 
Tight junction transmembrane complexes on the apical region of the 
intracellular cleft regulate and limit the paracellular flow of ions, solutes, lipids, and 
proteins. They prevent the diffusion of charged, large (>400 Da), and hydrophilic (>?8 
hydrogen bonds) molecules between the apical and basal sides of the cell and into the 
brain. They help establish an apical-basal polarity within the cell barrier [7]. Tight 
junctions are composed mainly of occludin and claudin proteins which are linked to 
the cytoskeleton via intracellular zonula occluden proteins (e.g., ZO-1/-2/-3). The near-
continuous formation and linkage of the claudin proteins between cells create a pore 
size of ~4.6 ? [99]. The different amino acid sequences within the first extracellular 
loop impose a charge selectivity on molecules that can diffuse past. [100]. Claudin-5, 
the most enriched tight junction protein at the BBB, is distinct to endothelial cells. The 
expression is regulated by VE-cadherin via VE-cadherin-mediated AKT activation and 
the exclusion of Forkhead box protein 01, a negative regulator of claudin-5 expression, 
20 
 
 
in the nucleus [101]. To further enhance the formation and integrity of the junction 
network, the nearby astrocytes and pericytes secret growth factors and morphogens 
(e.g., cyclic AMP, transforming growth factor-?, and bFGF), that regulate 
microvascular homeostasis [102, 103].  
It is increasingly clear that the presentation of junctional proteins (i.e., pattern, 
geometrical organization, edge-presentation) and cell morphology plays a vital role in 
regulating the permeability of cerebral endothelium [104-108]. As cell-cell junctions 
mature in vitro, VE-cadherin and ZO-1 reorganize from a serrated perpendicular 
geometry to a continuous strand linear to the cell edge [107-109]. Previous studies have 
shown junction maturity is dependent on extracellular matrix stiffness, and immature 
junctions are sites of drug penetration [108].  
2.2.3 Endothelial Cell Efflux Transporters 
In addition to restricting the entry of molecules into the brain via a physical 
barrier, the BBB is lined with ATP-binding cassette (ABC) efflux transporters that 
redirect drugs from the brain endothelial cells back into the bloodstream [110]. ABC 
transporters, localized to the luminal surface of endothelial cells, transport a wide range 
of substances against their concentration gradient via ATP hydrolysis [110]. ABC 
transporters are two homologous transmembrane proteins that adopt an inward-open 
confirmation with an exposed substrate-binding cavity. After a given substrate attaches 
to the binding domain and ATP binds to the nucleotide-binding domain, the protein 
shifts into an inward-closed confirmation, and the drug is transported out of the cell 
[111]. There are multiple types of ABC transporters present at the BBB that have 
overlapping substrate recognition [112]. 
21 
 
 
Drug efflux transporters comprise 10?15% of all proteins in the neurovascular 
unit [113]. Two of the most common ABC transporters at the BBB are ATP Binding 
Cassette Subfamily B Member 1 (ABCB1, also known as P-glycoprotein, P-gp) and 
ATP binding cassette subfamily G member 2 (ABCG2). Up-regulation of ABC 
transporters has traditionally been associated with multidrug cancer resistance of 
tumors. These transporters create a major hurdle when delivering drugs to brain cancers 
[88]. The development of ABCB1 and ABCG2 as therapeutic targets has been 
unsuccessful for many reasons including: (1) ABC transporters are found throughout 
the entire body and inhibition within normal tissue can result in additional side effects, 
(2) first generation inhibitors were not very potent, and patients were not screened for 
ABC expression before treatment, (3) pharmacokinetic interactions required 
chemotherapy dose reductions or resulted in the potential underdosing of patients, and 
(4) tumor cells and the BBB can contain multiple types of ABC transporters and the 
inhibition of one type can result in the over expression of others [114].  
For drug delivery to brain tumors to be effective, it must overcome the physical 
barrier created by tight junction proteins and the metabolic barrier created by ABC 
efflux transporters. This challenge highlights the necessity to develop a treatment 
strategy that can simultaneously modulate both proteins within the BBB.  
2.2.4 Drug Delivery to the Brain and Clinical Applications 
As a result of the physical and metabolic barriers present within the BBB, drug 
delivery to the brain is incredibly challenging. Numerous preclinical drug delivery 
systems have been engineered to overcome the BBB. These systems rely on disruption 
of BBB (paracellular route), enhanced transcellular transport, cell-based delivery 
22 
 
 
(transcellular route), or by passing the BBB via direct delivery to brain tissues (e.g., 
intrathecal, intraventricular, intranasal, and convection-enhanced delivery). 
Nanoparticle carriers have also been engineered with surface modifications that can aid 
in overcoming the BBB [8]. Most of these methods remain in preclinical studies due to 
low efficacy. 
Despite being the subject of studies for several decades, selective opening of 
the BBB junction network at the desired site without killing the brain endothelial cells 
remains challenging in the clinic. Several strategies have been developed to disrupt the 
tight junction network of the BBB, including the infusion of hyperosmolar solutions, 
biochemical disruption, focused ultrasound, and electromagnetic radiation  [8]. 
Nevertheless, these methods are not yet universally available in the clinic due to a 
number of limitations, such as low efficiency, invasiveness, lack of selectivity, or 
adverse events [8-11]. For example, clinical BBB opening with hypertonic solutions 
(e.g., arabinose, mannitol) lacks regional specificity and can induce a pronounced fluid 
shift [115, 116]. Existing site-specific methods (e.g., focused ultrasound [8, 117, 118], 
thermal therapy [119, 120], and radiation [121, 122]) can enhance drug delivery across 
the BBB but are often associated with increased risk of edema, hemorrhage, 
inflammation, or neurotoxicity. As a result, there is a clear need for a method that can 
dissociate the endothelial tight junction while sparing the endothelial cells. 
2.2.5 Relevance to Dissertation 
The intact BBB near the tumor-invaded brain regions limits chemotherapy 
effectiveness. Compared to the leaky BBB within the primary tumor, the BBB within 
the tumor-invaded brain regions stops almost all drugs from reaching and eradicating 
23 
 
 
the remaining infiltrative cells  [123]. One of the main focuses of this dissertation is to 
enhance drug delivery to the brain by repurposing a photochemistry-based cancer 
treatment modality and understanding the mechanisms behind BBB opening. Chapter 
4 will focus on understanding mechanistically how our selected treatment alters the 
cell-cell junctions in primary brain endothelial cells in vitro. We will further examine 
the drug delivery of a smaller molecule, dextran, and larger 100 nm liposomes across 
a monolayer of brain endothelial cells and determine how long the BBB remains ?open? 
after treatment. Chapter 7 will focus on engineering a drug delivery system and using 
a photochemistry-based treatment modality to open the BBB in rats. Chapter 8 will 
focus on developing a high throughput model for the screening of ABC inhibitors and 
present preliminary data on the photochemical modulation of efflux transporters. 
2.3 Photodynamic Therapy 
Light has been used in medicine for thousands of years. Ancient civilizations in 
Egypt, India, and China used light to treat skin diseases, including psoriasis and cancer 
[124]. In the 21st century, light and optics have significantly impacted modern 
medicine [125]. PDT is now a prominent tool being used in basic science and clinical 
practice to understand and manage disease [126-128]. PDT dates to the early 20th 
century in Germany when dyes and light were combined to treat skin cancer. Modern 
interest in PDT began in the 1960s with the discovery of the photosensitizer 
hematoporphyrin derivative [129]. Notably, Dr. Thomas Dougherty helped pioneer the 
field of photomedicine in the 1970s when he completed the first controlled clinical 
study in humans and was instrumental in the first FDA approval of a photosensitizer, 
Photofrin, for cancer treatment [130, 131]. 
24 
 
 
2.3.1 Fundamental Principles  
PDT is a photochemistry-based, non-thermal approach in which a non-
cytotoxic agent (i.e., photosensitizer) is energized by harmless light of an appropriate 
wavelength to produce cytotoxic agents. While there are multiple photochemical 
processes that an energized photosensitizer can undergo, this section will focus on the 
most predominant process, a type II photochemical reaction [13]. After absorbing a 
photon with a certain amount of energy, the photosensitizer's electrons will move into 
a single excited state. From here, the excited electrons can relax back to ground state 
or undergo intersystem crossing (i.e., altering their spin direction) and move into a 
triplet excited state. Now when the electrons relax from this longer-lived triplet state, 
an energy transfer to molecular oxygen can take place, resulting in the production of 
reactive oxygen species (ROS, e.g., 1O2, H2O2, O ?
-
2 , ?OH) [129]. For each photon 
absorbed by the photosensitizer, a single ROS may be produced. The predominant form 
of ROS produced is singlet oxygen (1O2), an oxidizing agent, which will be quantified 
in Chapters 3, 5, and 7. The excited state photosensitizer can also produce a 
fluorescence emission, instead of ROS, which will be discussed in section 2.3.3. After 
administration, and like chemotherapy, non-targeted photosensitizers will spread 
throughout the body. However, unlike chemotherapy, PDT treatment can be 
spatiotemporally controlled [18]. Generally, photosensitizers at doses used for PDT 
treatment are considered inert and won?t impact healthy tissues. After a set amount of 
time after photosensitizer administration and at the desired treatment site, light can be 
delivered at different powers and for different durations. When light delivery is 
stopped, the photochemical reaction is also immediately stopped. The generated 
25 
 
 
cytotoxic singlet oxygen is short-lived, with a half-life of 0.01?0.04 ?s, and will stay 
confined to the treatment site [132, 133]. 
 Numerous parameters impact the amount of singlet oxygen produced and PDT 
treatment outcomes. Depending on the amount produced, singlet oxygen can kill or 
modulate cells and tissues [23, 134], with threshold values required to induce tumor 
necrosis studied extensively by Dr. Theresa Busch [135]. Parameters, known as 
dosimetry (Figure 2.5) [132], that can impact PDT treatment include: 1. 
Photosensitizer type and tumor accumulation. Photosensitizing agents can have 
different backbones, such as porphyrins and chlorins. Not all photosensitizers will have 
the same singlet oxygen yield. Photosensitizers can be pro-drugs that are converted to 
an active form within a cell. Other photosensitizers require drug delivery vehicles 
which may alter their pharmacokinetics and biodistribution; 2. Photosensitizer 
activation wavelength and light dose parameters. Tissues have an extensive range of 
optical absorption and scattering coefficients, which can impact light penetration depth 
and photosensitizer activation. The laser power (W/cm2) and the total amount of light, 
fluence (J/cm2), a tissue receives, can impact photosensitizer photobleaching (i.e., the 
destruction of the ground state photosensitizer by ROS) and tissue oxygen 
concentrations; 3. Tumor oxygenation. A type II photochemical reaction requires 
ground state oxygen. Hypoxic tumors, where oxygen is limited, can diminish PDT 
efficacy [132].  
26 
 
 
Figure 2.5 Photodynamic Therapy Dosimetry Factors. Outlined are the interdependent 
factors, including photosensitizers, light, and oxygen, and the corresponding mechanisms that 
impact PDT treatment outcomes. Reprinted Wilson et al., Implicit and explicit dosimetry in 
photodynamic therapy: a New paradigm. Lasers Med Sci. 1997. Permission obtained from 
Springer Nature and Copyright Clearance Center?s RightsLink?.  
 
Light-activated photosensitizers produce singlet oxygen, an oxidizing agent, 
that inflicts direct damage to subcellular organelles (e.g., mitochondria, endoplasmic 
reticulum, lysosome), which can lead to cell death [129]. The mitochondria are one of 
the common sites for intracellular photosensitizer accumulation, including for the 
photosensitizer VP, which will be discussed further in Section 5.3.4 and throughout this 
work [136]. The production of singlet oxygen by light-activated photosensitizers can 
cause photodynamic disruption of the mitochondrial membrane (i.e., membrane 
depolarization), which triggers the release of cytochrome c, a potent initiator of 
27 
 
 
intrinsic apoptotic cell death [21]. After the release from the mitochondria, cytochrome 
c will interact with the apoptotic protease activating factor-1 in the cytosol, forming an 
apoptosome. Apoptosomes will go on to activate caspase-9 and eventually caspase-3, 
initiating a caspase cascade and cell death [137]. PDT has also been shown to 
photochemically degrade the anti-apoptotic protein Bcl-2 while leaving pro-apoptotic 
proteins Bad and Bax unaffected [138, 139]. This shifts the balance in target cells from 
an anti-apoptotic state to a more pro-apoptotic state [21]. These processes are known 
to bypass cell-death signaling pathways required for more traditional cancer treatments, 
such as chemotherapy. In addition, photosensitizers and PDT treatment does not result 
in the same side effects as chemotherapy [126, 134], allowing PDT treatment to be 
incorporated into the clinical workflow for many diseases. 
PDT typically involves the high optical energy (80-300 J/cm2) activation of 
photosensitizers [134, 140-143] and has historically been approved for the treatment of 
superficial diseases beneath the skin or in the lining of organs where a light source can 
reach [140]. With the advancement of fiber optic light conduits, there is now the clinical 
possibility for PDT to treat and permeabilize tumors that have grown deeply into the 
skin or other organs via intraoperative, endoscopic, or interstitial procedures [144]. 
Sections 2.3.2, 2.3.3, and 2.3.4 will now focus on different photosensitizers and PDT 
clinical applications. 
2.3.2 Photosensitizers 
Photosensitizers are non-toxic agents that can generate cytotoxic ROS upon 
optical irradiation with the appropriate wavelength of light. Tetrapyrrole structures 
such as porphyrins and chlorins are commonly studied and used in PDT treatments, 
28 
 
 
many of which are FDA approved. The aromatic structure of porphyrin, which contains 
four pyrrole rings and 18 ?- (or delocalized) electrons, makes it an ideal molecule for 
PDT applications. In general, photosensitizers used in clinical applications should have 
a strong absorption peak (i.e., q-band) in the red to the near-infrared spectral range, 
possess a large triplet quantum yield, have minimum dark toxicity effects (i.e., inert 
prior to light activation), and have relatively fast clearance [129, 145]. The optical 
therapeutic window for light-based treatments is considered 650-950 nm, where light 
has its maximum depth of penetration in tissue. Oxygenated hemoglobin strongly 
absorbs light below 600 nm, and water strongly absorbs light above 950 nm [146]. In 
addition, wavelengths longer than 850 nm do not provide enough energy to excite 
oxygen to its singlet state [129, 145].  
As mentioned previously, the first FDA-approved photosensitizer for PDT 
treatment was Photofrin in 1995. It remains one of the most widely used 
photosensitizers in the world [147]. However, Photofrin has a relatively low maximum 
absorption peak at 630 nm and an elimination half-life of 17 ? 4.3 days, leading to poor 
treatment outcomes and skin toxicity. Second-generation porphyrin-based 
photosensitizers have been developed to overcome these setbacks. Two 
photosensitizers discussed throughout this dissertation and in a direct comparison in 
Sections 3.3.1 and 3.3.2 are ALA-induced PpIX and VP (also known as benzoporphyrin 
derivative). ALA is a prodrug converted into its active form PpIX through the heme 
synthesis pathway. In healthy cells, PpIX is converted into the final product, heme, but 
in cancer cells, a lack of NADH results in low levels of ferrochelatase which results in 
a buildup of PpIX [148]. In 2017, the US FDA approved 5-aminolevulinic acid (5-
29 
 
 
ALA; Gleolan?) as an intraoperative imaging agent in patients with suspected high-
grade gliomas [149]. This will be discussed in more detail in Section 2.3.3.  
VP is a second-generation porphyrin-based photosensitizer. It has more ideal 
photochemical properties, including a 689 nm q-band for PDT treatment, a 24-hour 
elimination half-life, and an 86% singlet oxygen yield [150-153]. In 2001, the FDA 
approved the polydisperse liposomal formulation of VP, Visudyne?, for treating wet 
age?related macular degeneration. Free-form VP is not used clinically due to its 
inability to form sufficient hydrogen bonds with water. In an aqueous solution, free-
form VP will experience uncontrollable aggregation, resulting in a significantly 
reduced fluorescence emission signal, 1O2 production, and poor pharmacokinetics [21]. 
Visudyne? has been used worldwide to treat hundreds of thousands of patients, and 
evidence suggests treatment is safe, has minimal severe treatment side effects (e.g., 
only 1-4% of patients experience severe vision decrease), and can reduce the risk of 
visual loss in patients [154, 155]. For the treatment of age?related macular 
degeneration, Visudyne? is administered to patients intravenously, and about 15 
minutes later it is activated by a 689 nm laser to target and destroy the overgrown blood 
vessels within the eye [156].  Compared to ALA-induced PpIX, which requires 
intracellular conversion, VP can be activated on-demand at much shorter drug-light 
intervals. Since its approval, Visudyne? has been evaluated in numerous clinical trials, 
including for the treatment of refractory brain tumors (NCT00002647), pancreatic 
ductal adenocarcinoma (NCT03033225), and vertebral metastases (NCT02464761). 
While Visudyne? has shown promise in treating different cancers, the liposome can 
induce toxicity to normal tissues, trigger immunogenic responses, and reroute the 
30 
 
 
therapeutic payload to organs associated with the mononuclear phagocyte system 
(MPS) [6]. The immunological and toxicological consideration for liposome design 
will be discussed in depth within Chapter 6. The impact of lipids and other 
biomolecules on the photochemical properties of VP will be discussed in Chapter 5. 
Alternative drug delivery systems for VP will be discussed in Chapter 7.  
2.3.3 Fluorescence Guided Resection 
Besides generating cytotoxic ROS during PDT treatment, the singlet excited-
state photosensitizer can also emit a photon, which is helpful for fluorescence imaging 
and image-guided resection of tumors [157]. Surgical resection of tumors remains a 
first-line treatment modality for many cancer types, including GBM. The fluorescence 
signal produced by photosensitizers accumulated within a tumor can allow surgeons to 
better define tumor location and margins during the procedure. This enhanced surgical 
technique has increased the number of patients that can have a complete resection and 
improved safety by diminishing damage to normal tissue [158, 159]. Generally, 
fluorescence-guided resection (FGR) uses much lower laser powers and total fluence 
(> 5 mW/cm2, 0-20 J/cm2) and can also use blue light to excite the photosensitizers 
Soret band.  
Numerous photosensitizers have been used FGR imaging probes for a wide 
range of cancers. Notably, indocyanine green is approved for laparoscopic surgery, 
methylene blue is used to visualize ureters during colorectal procedures and map 
sentinel lymph nodes during breast cancer surgeries, and the cetuximab-IRDye800CW 
was recently approved in Japan for use in head and neck cancers [158]. One particularly 
successful photosensitizer for the FGR of GBM is ALA?induced PpIX [149]. In 2017, 
31 
 
 
the FDA approved 5-ALA (Gleolan?) as an intraoperative imaging agent in patients 
with suspected high-grade gliomas [149]. Upon excitation with ultraviolet/blue light 
(375-440 nm), gliomas that contain the 5-ALA-induced protoporphyrin IX (PpIX) emit 
a violet-red fluorescence, which the naked eye can easily visualize (i.e., without filters) 
for resection [160]. Fluorescence-guided brain tumor surgery is routinely applied in 
some clinical centers, whereas the combination of FGR and PDT is under active 
preclinical and clinical studies. While this FDA-approved surgical technique has shown 
to increase the mean patient survival by 3 months compared to standard white light 
surgery [159], FGR and ALA-mediated PDT have not yet achieved long-term survival 
[161]. Fluorescence imaging of PpIX and VP will be discussed further in Sections 3.3.2 
and 5.3.4. 
Figure 2.6 ALA mediated Fluorescence Guided Resection of GBM. (a) White light surgery 
shows a relatively homogenous tumor appearance. (b) ALA induced PpIX fluorescence under 
blue excitation light helps identify the cancerous lesion. Reprinted Hadjipanayis, et al., What 
is the Surgical Benefit of Utilizing 5-ALA for Fluorescence-Guided Surgery of Malignant 
Gliomas? Neurosurgery. 2015 Nov; 77(5): 663?673. Permission obtained from Wolters 
Kluwer and Copyright Clearance Center?s RightsLink?. 
 
2.3.4 Clinical Applications 
PDT has shown promise in treating different ailments. FDA approvals include 
treatment of head and neck cancers [162], esophageal cancer [163], non-small cell lung 
32 
 
 
cancer [164], and advanced cutaneous T-cell lymphoma [165]. It is also approved in 
Mexico for prostate cancer [166] and Japan for unresectable locally advanced or 
recurrent head and neck cancer (NCT03769506). The remaining sections and chapters 
will focus on using ALA-induced PpIX and VP to treat GBM. As previously 
mentioned, ALA is a prodrug administered as an oral solution to patients. Alternatively, 
in aqueous solutions, free-form VP will experience uncontrollable aggregation, 
resulting in complete self-quenching, minimum singlet oxygen production, and poor 
pharmacokinetics [21]. Therefore, for effective patient delivery, VP must be 
solubilized or incorporated into a drug delivery vehicle. The current gold standard for 
delivering VP to patients is in its phospholipid-associated form, Visudyne?. Adding 
5ml of water to the freeze-dried lipids and VP followed by vigorous shaking results in 
a self-assembled polydisperse liposome that has an average diameter of 524.8 ? 547.5 
nm and a VP concentration of ~2.8mM [167]. Visudyne? is being tested in several 
clinical trials evaluating PDT for pancreatic ductal adenocarcinoma (NCT03033225) 
[168], recurrent and metastatic prostate cancer (NCT03067051, NCT02939274), breast 
neoplasms (NCT02872064), and pleural malignancies (NCT02702700). Surgery 
immediately followed by PDT has been studied in the clinic for brain tumors 
(NCT03048240, NCT03897491, NCT00003788) [143, 169]. In fact, a single 
adjunctive dose of PDT has been shown to add up to 18 months to the lifespan of 
patients with GBM [12]. 
2.3.5 Photodynamic Priming 
Targeting the tumor microenvironment, including the vasculature, could 
provide additional opportunities to modulate tumor physiology and enhance the 
33 
 
 
delivery of therapeutic agents for improved treatment outcomes. Previous studies have 
demonstrated the ability of PDT to permeabilize tumor vasculature and the BBB for 
enhanced intratumoral drug delivery [14-20]. However, despite the promising 
preclinical results using PDT to permeabilize the BBB and treat brain tumors, the side 
effects, including edema, neurotoxicity, and even animal death, remain a major concern 
in the clinic [15, 170-172]. A study by Hirschberg and colleagues in 2007 showed that 
the human equivalent dose of the photosensitizer ALA (125 mg/kg) and a light dose of 
26 J could open the BBB for up to 3 days, but also resulted in the death of 25% of 
tumor-bearing rats. To address this issue, our lab has introduced priming which 
operates at low doses of optical energy and photosensitizer (i.e., in vitro dose is <1 ?M 
? J/cm2) to modulate biomolecular targets without killing the organism. Leveraging 
subtherapeutic doses of PDT can also impact tumor immune response and overcome 
drug resistance by modulating efflux transporters. 
Traditional PDT uses light doses of 80-300 J/cm2 to induce cancer cell killing. 
Traditional PDT also uses large photosensitizer doses (e.g., 2.0 mg/kg VP in mice, the 
human equivalent dose [173]) and a longer drug-light interval to ensure the 
photosensitizer accumulates within both the vasculature and the tumor. Alternatively, 
PDP uses light doses in the range of 10-80 J/cm2 to modulate the vasculature without 
damaging the surrounding tissues. PDP uses smaller photosensitizer doses (e.g., 0.25 
mg/kg in mice, ? the human equivalent dose [19])  and a shorter drug-light interval, 
usually between 15-90 minutes, to ensure the photosensitizer is confined to the 
vasculature [174]. Encouraging results have been obtained with PDP for pancreatic and 
brain tumors. Chapter 4 will explore how PDP can be used to disassociate the 
34 
 
 
endothelium of the BBB that has an exceptionally high degree of junctional tightness 
without damaging the surrounding healthy brain. 
2.3.6 Relevance to Dissertation 
As is evident from the numerous clinical trials (NCT03048240, NCT03897491, 
NCT00003788, NCT01966809, NCT00002647, NCT01682746), the photochemistry-
based treatment modality, PDT, has shown promise in treating GBM. PDT has also 
shown promise in opening the tumor vasculature and enhancing intratumoral drug 
delivery. Our lab has recently shown that sublethal PDT improves vascular 
permeability and enhances ONIVYDE? (Liposomal irinotecan) extravasation into 
pancreatic tumor parenchyma by up to 11-fold without damaging the surrounding 
healthy pancreas [175]. This dissertation will expand our knowledge and focus on how 
PDT can be used to enhance drug delivery to the brain and treat brain tumors.  
Throughout this dissertation, photosensitizers, PDT, and PDP will be examined 
for the treatment of GBM. Chapter 3 will focus on light penetration depth in the rodent 
brain. A direct comparison between PpIX and VP will be performed. Chapter 4 will 
focus on the meninism behind PDP-induced BBB opening to enhance drug delivery to 
residual GBM cells. Chapter 5 will focus on improving VP photochemical properties 
via biomolecule-VP conjugates. Lastly, Chapter 7 will focus on a pure-drug carrier-
free nanoparticle of VP for GBM treatment.  
 
 
 
 
35 
 
 
3. Chapter 3. Photodynamic Effect Depth in the 
Rodent Brain 1 
3.1 Introduction 
Light and optics have had significant influence on modern medicine [125]. 
PDT, a major output of this work, is now being used extensively in basic science and 
clinical practice to understand and manage disease [126-128]. FGR and PDT using 
different photosensitizers (e.g., VP, 5-ALA, Photofrin?, temoporfin, hematoporphyrin 
derivative) have been evaluated clinically for patients with brain cancer (e.g., 
NCT03048240, NCT03897491, NCT00003788, NCT01966809, NCT00002647, 
NCT01682746). The pro-drug photosensitizer, ALA (Gleolan?) was approved in 2017 
as an intraoperative imaging agent in patients with high-grade gliomas [149]. In 
addition, a single adjunctive dose of PDT has been shown to add as much as 18 months 
to the lifespan of GBM patients [12]. In a small-scale study, the combination of FGS 
and PDT increased the mean survival of GBM patients from 24.6 to 52.8 months, 
compared to FGS alone [176]. However, long-term survival using FGS and/or PDT has 
not been achieved. It is well recognized that limited light penetration in brain tissues, 
and the poor selectivity and delivery of the photosensitizers impacts treatment 
outcomes.  
Tissue optical properties influence light delivery and impact PDT outcomes 
[177]. Red light penetration depth is typically 0.1-0.3 cm in tissues [125]. This 
                                                 
1 This chapter was adapted from Inglut C.T., Gaitain B., et al. ?Predictors and Limitations of 
the Penetration Depth of Photodynamic Effects in the Rodent Brain?. Photochem 
Photobiol.2020 Mar;96(2):301-309. Permission obtained from John Wiley and Sons to use this 
material in this dissertation. B.G. was a co-author. He co-wrote several sections in this chapter, 
and helped perform and analyze the experiments for Figures 3.4. 
36 
 
 
penetration depth is defined as the depth at which the incident optical energy drops to 
1/e (~37%) [125]. A number of ex vivo and in silico studies have already shown that 
the remaining 37% of photons can reach up to 2-5 cm deep in brain tissues [178-184]. 
A series of studies by Wilson, Perria, and Cheng have shown that 630 nm light 
activation of hematoporphyrin derivative induces tumor necrosis and cell killing at 0.8-
1.5 cm deep within the brain [185-187]. On the other hand, Olzowy et al. demonstrated 
that 630 nm light activation (200 mW/cm2) of 5-ALA-induced PpIX induces cerebral 
tissue damage only up to 0.4 cm deep [188]. Based on prior work showing that 690 nm 
light can penetrate approximately 40% further than 633 nm light in bladder tissue [189], 
here, we test if VP could be activated by 690 nm light at further depths in the brain, 
compared to using 635 nm light-activated PpIX. 
Both VP and PpIX are hydrophobic molecules and tend to aggregate in 
biological environments, leading to low bioavailability, heterogeneous 
biodistributions, and reduced PDT efficacy. Nanoliposomes, composed of a 
hydrophobic lipid bilayer and an aqueous core, are the most studied and clinically used 
photosensitizer delivery vehicles [190]. In 2001, a polydisperse liposomal formulation 
of VP (Visudyne?, 0.2-1 ?m in diameter) was approved by the FDA for treatment of 
patients with wet age-related macular degeneration [145, 191]. A clinical study using 
Visudyne?-PDT to treat patients with refractory brain tumors is also taking place 
(NCT00002647) [192]. While PpIX production in tumors is commonly derived from 
the oral or systemic administration of 5-ALA prodrug [149], there has been a 
resurgence of interest in PpIX delivery using different nanoformulations to, in part, 
bypass the heterogeneous and low conversion of 5-ALA to PpIX observed in some 
37 
 
 
tumors [193-195]. Based on these advances, here, we synthesized monodisperse 
Liposomal formulations of VP (Liposomal VP) and PpIX (Liposomal PpIX). Using 
these photosensitizing liposomes, we examined the limits of PDT as a function of tissue 
depth and light fluences in ex vivo rat brains, by monitoring the changes in 
photosensitizer photobleaching and singlet oxygen (1O2) production. This chapter will 
give us insights into which photosensitizer is more advantageous for the targeting of 
microscopic GBM infiltrates that can be up to 2cm away from the primary tumor.  
3.2 Materials and Methods 
3.2.1 Synthesis and Characterization of Liposomal VP and Liposomal PpIX 
Liposomes containing VP or PpIX within its bilayer (Liposomal VP and PpIX) 
were synthesized via freeze-thaw extrusion as described previously [19, 175, 196-198]. 
Briefly, lipids, dipalmitoylphosphatidylcholine (DPPC), cholesterol, and 
distearoylphosphatidylethanolamine-methoxy polyethylene glycol (DSPE-PEG), and 
1,2-dioleoyl-3-trimethylammonium-propane (DOTAP) (Avanti Polar Lipids), were co-
dissolved with VP (50 nmoles, U.S. Pharmacopeial) at 0.15 mol% VP-to-lipid ratio or 
PpIX (60 nmoles) at 0.18 mol% PpIX-to-lipid ratio. Chloroform was removed via 
rotary evaporator overnight in order to create a thin film, which was rehydrated with 1 
mL of deionized water. The lipid suspension was subjected to freeze-thaw cycles (4-
45?C). The dispersions were extruded through polycarbonate membranes (0.1 ?m pore 
size) at 42?C to form unilaminar vesicles. The final concentration of the dispersions 
was estimated by absorbance using the established molar extinction coefficient of VP 
in DMSO (~34,895 M-1cm-1 at 687 nm) or PpIX in chloroform (~171,000 M-1cm-1 at 
38 
 
 
407 nm) or DMSO (~170,000 M-1cm-1 at 407 nm). The fluorescence properties 
Liposomal VP (Excitation/Emission: 435/650-750 nm) and Liposomal PpIX 
(Excitation/Emission: 405/600-800 nm) was determined using a multi-mode 
microplate reader (Synergy Neo2, BioTek). Particle size and zeta potential were 
measured on the Nanobrook (Brookhaven). Photosensitizers were diluted to 5 ?M in 
phosphate buffered saline (PBS) for all optical-based characterization. The production 
of singlet oxygen from photoactivated VP and PpIX was monitored indirectly using the 
probe, Singlet Oxygen Sensor Green (SOSG). Briefly, 0.5 ?M Liposomal VP or 
Liposomal PpIX were mixed with 25 ?M SOSG (Thermo Fisher Scientific) in PBS. 
Samples were irradiated with 635 nm light (100 mW/cm2) for Liposomal PpIX or 690 
nm light (100 mW/cm2) for Liposomal VP, at different fluences of 0-80 J/cm2 (ML6600 
series laser module, Modulight). The fluorescence intensity of the cleaved SOSG probe 
(Excitation/Emission: 504/525 nm) was acquired using a microplate reader prior to and 
following irradiation. 
3.2.2 Photobleaching of Photosensitizers in Ex Vivo Rat Brains 
The activation of VP and PpIX was measured via photobleaching in Sprague 
Dawley rat brains (Innovative Research). Glass capillaries (300 ?m in diameter; 1 cm 
in length) containing 5 ?M of photosensitizer were placed in the brains at varying 
depths (0-2 cm). The brain tissues were exposed to 635 nm red light for Liposomal 
PpIX or 690 nm Near-infrared light for Liposomal VP using a diode laser (ML6600, 
Modulight). A range of fluences from 0-80 J/cm2 (100 mW/cm2) were examined. After 
PDT, capillaries were removed, and the fluorescence emission from Liposomal VP (or 
39 
 
 
Liposomal PpIX) was measured using a reflectance fluorescent microscope equipped 
with a 375 nm laser diode (L375P70MLD, Thorlabs). 
3.3 Results and Discussion 
3.3.1 Synthesis and Characterization of Liposomal VP and Liposomal PpIX 
Liposomal VP and Liposomal PpIX (Liposomal PpIX) were reproducibly 
synthesized via the freeze-thaw extrusion method [19, 175, 196, 197]. A lipid 
composition of dipalmitoylphosphatidylcholine (DPPC), cholesterol, and 
distearoylphosphatidylethanolamine-methoxy polyethylene glycol (DSPE-PEG) at a 
20:10:1 molar ratio was used based on the clinically approved PEGylated Liposomal 
formulations [199, 200]. Here, both Liposomal VP and Liposomal PpIX were grafted 
with 3 mol% of PEG and formed in the size range of ~135-150 nm with a narrow size 
distribution (polydispersity index, PdI < 0.1). Cationic 1,2-dioleoyl-3-
trimethylammonium-propane (DOTAP, 8 mol%) was introduced to the lipid 
composition to provide a zeta-potential of +10-12 mV (Table 3.1). Photosensitizers 
(VP or PpIX) were adsorbed in the liposomal lipid-bilayer via hydrophobic and ionic 
interactions [201]. Our stability results suggest that 1-4 months of dark storage at 4?C 
and in PBS did not affect the overall size or monodispersity (PdI <0.1) of Liposomal 
VP (139.7?11.8 nm in diameter) and Liposomal PpIX (150.9?6.6 nm in diameter) 
40 
 
 
 
Table 3.1 Liposomal VP and PpIX. Overview of the characterization of Liposomal VP and 
Liposomal PpIX detailing their respective hydrodynamic diameter, polydispersity index, and 
zeta potential. Values are presented as mean?standard deviation. N ? 3, where N equals the 
amount of samples synthesized.  
Parameters L-VP L-PpIX
 Hydrodynamic diameter (nm) 135.7 ? 8.0 153.1 ? 4.9
Polydispersity index (PdI) 0.06 ? 0.02 0.04 ? 0.02
Zeta potential (mV) 10.1 ? 1.2 12.4 ? 1.8
 
While liposomes can maintain the stability and photoactivity of 
photosensitizers in biologically relevant media for effective PDT, hydrophobic 
photosensitizers in the lipid bilayer of liposomes are prone to quenching [201-203]. To 
test if the unquenched photosensitizers (VP or PpIX) are stabilized in the liposomes 
and could be photo-activated for PDT, we evaluated the stability, quenching factor, and 
singlet oxygen (1O2) production of each photosensitizer in both free and liposomal 
forms, with or without light activation. Changes in photosensitizer absorbance and 
fluorescence were monitoredand in PBS (Figure 3.1a, b). In Figure 1a, the absorbance 
(blue dotted line) and fluorescence (red dotted line) spectra for free VP in PBS were 
significantly lower than that of Liposomal VP (solid lines) in PBS, indicating the rapid 
aggregation of free VP molecules under physiological conditions without being 
encapsulated in a liposomal formulation. Similarly, ?recovery? of the absorbance (blue 
dotted line) and fluorescence (red dotted line) spectra of PpIX photosensitizers in PBS 
was made possible by the liposomal formulation (Figure 3.1b). These results suggest 
that the unquenched photosensitizers (VP or PpIX) are stabilized in the liposomes and 
could be activated by light for PDT, even without the release of photosensitizers from 
the liposomes. 
41 
 
 
Light activation of photosensitizers results predominately in a type II reaction 
which produces highly reactive 1O2 [23]. The 
1O2 quantum yield of VP (??= 0.86) is 
significantly higher than that of PpIX (??= 0.56) [150-153]. Using the SOSG probe, 
we indirectly evaluate if 690 nm light activation of VP could produce 1O2 more 
efficiently than 635 nm photo activated PpIX in PBS. Light activation of VP generated 
a markedly higher SOSG fluorescence intensity than that generated by PpIX, in an 
energy dependent manner (Figure 3.2a). The SOSG fluorescence signal began to 
plateau at 60-80 J/cm2. It is speculated that an insufficient supply of ground state 
oxygen or available molecular probe lowered the SOSG signal at 80 J/cm2. These 
results indicate Liposomal VP can produce more 1O2 than Liposomal PpIX upon light 
activation under our experimental conditions.  
 
Figure 3.1 Absorbance and fluorescence spectra of VP and PpIX (a) Free-form VP fully has a 
strong absorbance peak at 435 nm (soret band) and 690 nm (Q band). Typically for PDT, VP is 
activated at 690. VP fluoruence emission has a peak in the near-infrared range (?max at ~700 nm).  
Free VP is hydrophobic and readily aggregates in PBS, as demonstrated by the decrease in 
magnitude of the absorbance spectrum, resulting in fluorescence quenching of VP with minimal 
photoactivity (< 2%). Encapsualting VP in a Liposomal formulation restores its fluorescence and 
photoactivity (35.8?0.2%) in PBS. (b) Free-form PpIX fully shows a strong absorbance peak at 
405 nm (soret band), with multiple Q bands in the green to red range. Typically, PpIX is activated 
by 635 nm light or 375-440 nm light for PDT or FGS, respectively. The fluorescence signal emitted 
by PpIX is in the red/ near-infrared range between 635 and 700 nm. Likewise, free-form PpIX 
aggregates in PBS, leading to a quenched state that is no longer activatable at 635 nm. Similarly, 
the encapsulation of PpIX within a Liposomal formulation restores its fluorescence and 
photoactivity up to 82.8?0.1%. N ? 6, where N equals the number of wells conducted over 3 trials. 
42 
 
 
3.3.2 Evaluation of the Tissue-depth Dependent Photodynamic Effects of VP and 
PpIX 
Photobleaching is a process in which a photosensitizer (or fluorophore) 
permanently loses the ability to fluoresce and/or undergo type I/II reactions due to 
photochemical damage and covalent modification of the photosensitizer. Measuring  
the loss of fluorescence as the photosensitizer is photobleached is perhaps the most well 
studied and easily accessible tools for PDT monitoring and dosimetry in preclinical 
(Figure 3.2b) and clinical settings [204-207]. For example, Tyrrell et al. showed 
patients lesions with significantly higher levels of PpIX photobleaching post PDT had 
complete clearance at 3 months [208]. Here, we examined the penetration depth of the 
photodynamic effects in rat brains by monitoring the loss of fluorescence, as the 
photosensitizer is photobleached, after surface light illumination. Briefly, capillaries 
filled with 5 ?M of Liposomal VP or Liposomal PpIX, were inserted into the ex vivo 
rat brains at varying heights followed by surface illumination using appropriate 
wavelengths (i.e. 690 nm for Liposomal VP; 635 nm for Liposomal PpIX) (Figure 
3.3). After light activation, the total fluorescent signal was compared to a negative 
Figure 3.2 Singlet Oxygen Production and Photobleaching of Liposomal VP and 
Liposomal PpIX. (a) SOSG fluorescence emission and (b) photobleaching percentage of 
Liposomal VP and Liposomal PpIX with respect to increasing fluences of light at 690 nm and 
635 nm, respectively. N ? 6, where N equals the number of wells assessed over 3 trials. 
43 
 
 
control to determine the degree of photobleaching. A non-linear decrease in the 
percentage of Liposomal VP (or Liposomal PpIX) photobleaching was observed with 
increased tissue depth and reduced fluence (Figure 3.4a,b). Up to 10% photobleaching 
of Liposomal VP was observed at 1.4-2 cm (Figure 3.4a,c), while negligible (0-2%) 
photobleaching of Liposomal PpIX was observed at 1 cm deep in brain tissue (Figure 
3.4b,c). This result appears to be similar to previous findings [209, 210]. Dereski and 
colleagues demonstrated that Photofrin?, upon 632 nm light activation, induces a 
necrotic zone ~0.35 cm deep within male Fisher rat brains [209]. A clinical study 
showed talaporfin, which is activated at a longer wavelength of 664 nm, could induce 
PDT damage (based on demyelination) up to 0.8 cm in a patient with recurrent glioma 
[210]. It has also been shown that illumination geometry impacts the depth of brain 
tissue damage induced by PDT. Specifically, using hematoporphyrin derivative-based 
PDT, Perria et al. found that the depth of necrosis with surface illumination was 1.5 cm 
deep, compared with the 0.8 cm radius of cell killing found by Cheng et al. using an 
implanted optical fiber [185-187]. Further studies comparing surface illumination with 
illumination using implanted optical fibers are warranted. 
  
 
 
 
 
 
 
44 
 
 
Figure 3.3 Experimental setup for examining the limit of the photodynamic effect. Glass 
capillaries containing Liposomal VP or Liposomal PpIX (5 ?M) were inserted at various 
depths (Z) in rat brains and exposed to 690 nm or 635 nm light (surface illumination) 
The degree of photobleaching across the different energies (40 and 80 J/cm2) 
converge as depth increases (Figure 3.4a,b) since the loss of light intensity is non-
linear. Light penetration can be modeled using the simple 1-dimensional (1-D) equation 
?(?) = ?????? [211], where ?(?) is the fluence rate (W/cm2) dependent on depth 
(cm), z. E is the irradiance delivered on the surface of the tissue (W/cm2), k is 
backscattered light that augments irradiance on the surface of the tissue, and ? is the 
attenuation coefficient (cm-1). The amount of energy lost is modeled as an exponential 
decay (???). Therefore, it is expected that the depth of photodynamic effects would be 
proportional to ?(?), and the depth versus degree of photobleaching should also have 
an exponential decay. However, due to the specular reflectance when the light hits the 
brain, and the boundary condition of the air-brain barrier, the trend shows a plateau 
followed by an exponential decay (Figure 3.4a,b). Yet, based on the model set up, the 
light initially penetrating the brain at very shallow depths should have a fluence rate 
slightly greater than the initial irradiance, and photobleaching should be greater [212-
45 
 
 
214]. Light scattering in tissue is anisotropic (i.e., has a g factor of 0.9 on a -1 to 1 
scale), meaning most light will forward scattering and a small amount of light is 
scattered back [215]. There is also a possibility of some backscattered light that hits the 
air-brain barrier to be reflected back into the brain again. This additional light scattering 
will increase the fluence rate as depth increases. However, based to the depth intervals 
taken, this phenomenon cannot be seen (Figure 4a,b). 
 
Figure 3.4 Depth of Photosensitizer Activation within Brain Tissue Determined via 
Degree of Photobleaching.  Photobleaching of (a) Liposomal VP and (b) Liposomal PpIX 
were evaluated at different depths and fluences. (c) At 80 J/cm2, 27% Liposomal VP 
photobleaching was observed 1 cm deep in tissue, in contrast to only 0-2% photobleaching of 
Liposomal PpIX. N ? 3. 
 
The fact that PpIX demonstrated a shallower penetration depth than VP is not 
all that surprising. The main reason comes down to the activation wavelength. VP was 
activated using a 690 nm wavelength and PpIX was activated with a 630 nm 
46 
 
 
wavelength. Although this is only a 60 nm shift, there is an appreciable drop in 
absorption by up to 40-50% [216-218]. This led to the 690 nm light penetrating double 
the depth of 630 nm light [219]. The depth of treatment is described with the equation 
???
??? = ?ln?( ) [213]. Zrx is the depth penetration of the photodynamic effect, ? is ???
optical penetration depth (cm), t is the exposure time (s), and Hth is the minimum 
amount of energy needed to make the photodynamic effect on the tissue (W*s/cm2). 
Looking at this equation, increasing the E or t increases the depth of the effect by ln(Et), 
while increasing the wavelength would increase the optical penetration depth, which as 
a result would increase the penetration depth of the photodynamic effect in a 
proportional manner. 
While Liposomal VP has greater potential for deeper activation and larger 
production of 1O2 in tissue, ALA-induced PpIX is a valuable tool for the FGS of 
gliomas.  A digital comparison of the two capillary tubes (Figure 3.5) filled with 
Liposomal VP (top) and Liposomal PpIX (bottom) reveal PpIX can be seen easier 
compared to VP. The capillaries were placed at similar heights, 0.04-0.05 cm below 
Figure 3.5 Visualization of the Fluorescent Emission of Liposomal VP and Liposomal 
PpIX when Excited at 375 nm. (a) Capillaries filled with the photosensitizers (Liposomal 
VP or PpIX, 5 ?M) were excited with 375 nm and captured with a phone camera (OnePlus 5). 
PpIX?s fluorescent signal can be easily visualized by the naked eye. (b) Capillaries were 
inserted into the brain and were excited to examine if the fluorescent signal could be visualized 
through brain tissue. The emission signal from PpIX could be seen when the capillary was 
inserted ~0.04-0.05 cm deep in tissue (black arrow). 
47 
 
 
the surface of the brain and excited with 375 nm light, a similar wavelength of light 
used for FGS. The higher fluorescence quantum yield of PpIX (?F= 0.155) and its 
fluorescence peak at 630 nm make it easier for the naked eye to visualize. VP fluoresces 
at 700 nm and has a lower fluorescence quantum yield (?F= 0.051) [150-153]. Even if 
the same number of photons were being emitted, the emitted 630 nm light is seen with 
10 times greater sensitivity by the human eye than the 690-710 nm light [220]. 
In summary, the results show that the liposomal formulations improve the 
stability and photoactivity of VPand PpIX in PBS. Near-infrared light (690 nm) 
activation of Liposomal VP induces 1O2 production, which causes photobleaching of 
VP up to 2 cm deep in brain tissues. In contrast, no photobleaching was observed 
beyond 1 cm deep in brain tissues upon red light (635 nm) activation of Liposomal 
PpIX. The enhanced penetration depth of Liposomal VP-mediated PDT effects in the 
brain tissue paired with the use of PpIX for tumor visualization opens the door to 
developing and optimizing the combination of FGS and PDT for recurrent brain 
tumors, which typically occur within 1-2 centimeters of the original tumor [44, 45, 
221]. Characterization and optimization of Liposomal PpIX based FGS and Liposomal 
VP-mediated PDT in ex vivo brain tissues will pave the way towards investigating these 
new concepts in animals and eventually in patients. 
 
 
48 
 
 
4. Chapter 4. Understanding the Effects of Photodynamic 
Priming on Endothelial Cell?Cell Junctions for 
Enhancing Drug Delivery2 
4.1 Introduction 
As described in Section 2.2, the unique capillaries within the brain, known as 
the BBB, limit the diffusion of large or small compounds from the bloodstream into 
the brain. This specialized neurovascular unit, composed of endothelial cells, neural 
cells (e.g., astrocytes, pericytes, neurons) and basement membrane, regulates brain 
homeostasis and protects the central nervous system [81-83]. The cell-to-cell junctions 
that join the brain endothelial cells are composed of a wide variety of components, such 
as vascular endothelial-cadherin (VE-cadherin) adherens junction protein, zonula 
occludens (ZO) scaffold proteins, and claudin-5 tight junction protein. The tight 
junction transmembrane complexes located on the apical region of endothelial cells 
prevent the transport of hydrophilic, highly charged, and/or large (>400 Da) molecules 
[7]. In addition, astrocytes and pericytes secret growth factors and morphogens (e.g., 
cyclic AMP) to further enhance the formation and integrity of the junction network 
[102, 103]. The endothelial cell-cell junction at the BBB is one of the key factors that 
limit drug transport to the brain [87, 95]. Despite being the subject of studies for several 
decades, selective opening of the BBB junction network at the desired site without 
killing the brain endothelial cells remains a challenging task in the clinic. 
                                                 
2 This chapter was adapted from Inglut C.T., Gray K.M., et al., ?Photodynamic Priming 
Modulates Endothelial Cell?Cell Junction Phenotype for Light-activated Remote Control of 
Drug Delivery?. IEEE J Sel Top Quantum Electron. Jul-Aug 2021;27(4):7200311. Permission 
obtained from IEEE to use this material in this dissertation. K.M.G helped perform and analyze 
the experiments for Figures 4.2, 4.3, and 4.4. 
49 
 
 
A number of strategies have been developed to disrupt the tight junction 
network of the BBB, including the infusion of hyperosmolar solutions, biochemical 
disruption, focused ultrasound, and electromagnetic radiation [8]. Nevertheless, these 
methods are not yet universally available in the clinic due to a number of limitations, 
such as low efficiency, invasiveness, lack of selectivity, or adverse events [8-11]. For 
example, clinical BBB opening with hypertonic solutions (e.g., arabinose, mannitol) 
lacks regional specificity and can induce a pronounced fluid shift [115, 116]. Existing 
site-specific methods (e.g., focused ultrasound [8, 117, 118], thermal therapy [119, 
120], and radiation [121, 122]) can enhance drug delivery across the BBB but are often 
associated with increased risk of edema, hemorrhage, inflammation, or neurotoxicity. 
As a result, there is a clear need for a method that can dissociate the endothelial tight 
junction while sparing the endothelial cells. 
PDT is a photochemistry-based treatment modality that involves the light 
activation of non-toxic photosensitizers or fluorescent dyes. High optical energy (e.g., 
80-300 J/cm2) activation of photosensitizers is used to generate reactive oxygen species 
that can damage the vasculature and kills cells [175]. In fact, surgery immediately 
followed by PDT has already been studied in the clinic for brain tumors 
(NCT03048240, NCT03897491, NCT00003788) [143, 169]. A single adjunctive dose 
of PDT has been shown to add up to 18 months to the lifespan of patients with GBM 
[12]. Despite the promising preclinical and clinical results using PDT to permeabilize 
the BBB and treat brain tumors, the side effects of PDT, including edema and 
neurotoxicity, remain a major concern in the clinic [15, 170-172].To address this issue, 
we have introduced PDP which operates at low doses of optical energy and 
50 
 
 
photosensitizer (i.e., < 1 ?M ? J/cm2) to modulate biomolecular targets without killing 
the organism [175]. We have shown that PDP improves vascular permeability and 
enhances ONIVYDE? (Liposomal irinotecan) extravasation into pancreatic tumor 
parenchyma by up to 11-fold without damaging surrounding healthy pancreas [175]. A 
combination of PDP and ONIVYDE? inhibited tumor growth, reduced metastasis, and 
doubled the median progression-free survival in pancreatic cancer mouse models [19, 
175, 222]. While encouraging results have been obtained with PDP for pancreatic 
cancer treatment, there is work to be done to ensure that PDP can be used to disassociate 
the endothelium of the BBB that has a particularly high degree of junctional tightness. 
It is increasingly clear that the presentation of junctional protein (i.e., pattern, 
geometrical organization, edge-presentation) and cell morphology plays an important 
role in regulating the permeability of cerebral endothelium [104-108]. Despite that 
information, the impact of sub-cytotoxic photochemistry on the presentation of 
junctional protein remains relatively understudied, presumably due, in part, to the lack 
of reliable quantitative analysis routines. Here, for the first time, we utilized a novel 
Junction Analyzer Program (JAnaP) [105, 223] to quantify junction phenotype in 
response to PDP at various light energy densities using human brain microvascular 
endothelial cells (HBMECs). Images of immuno-stained ZO-1 and VE-cadherin were 
analyzed using the JAnaP to calculate the percent of the cell perimeter presenting 
continuous, punctate, or perpendicular junctions before and after PDP. Furthermore, a 
transwell permeability assay was used to demonstrate that BBB permeability to 
fluorescein isothiocyanate?dextran (FITC-dextran) or rhodamine-loaded liposomes 
(rhodamine-liposome) can be precisely controlled via light activation. This chapter will 
51 
 
 
give insights into how PDP modulates cell-cell junctions and how it can be used to 
enhance drug delivery past the endothelial cell barrier.  
4.2 Methods 
4.2.1 Cell Culture 
Primary HBMECs were purchased from Cell Systems (ACBRI 376) and 
cultured as previously described [108]. Briefly, cells were cultured in flasks coated with 
0.1% gelatin and grown in RPMI-1640 medium supplemented with 20% FBS, 1% 
Pen/Strep, 2 mM L-glutamine (Thermo Fisher Scientific, Waltham, MA), 30 ?g/mL 
endothelial cell growth supplement (Millipore Sigma, Burlington, MA), and 100 
?g/mL heparin (Millipore Sigma) at 37 ?C, with 5% CO2. Cells were used for 
experiments within 7-10 passages. Cells arrived mycoplasma-free and were tested and 
found to be mycoplasma-free approximately 6 months later using the MycoAlert PLUS 
Mycoplasma Detection Kit (Lonza, LT07-701). 
4.2.2 Substrate Coating and Experimental Conditions 
On day 1 of experiments, glass bottom 24-well plates (662892, Greiner Bio-
One, Kremsm?nster, Austria) or transwell inserts (24 well format, 1.0 ?m pore size, 
Falcon, Abilene, TX) were respectively coated with 175 ?L or 35 ?L of 100 ?g/mL 
fibronectin (F2006, Sigma Aldrich, St. Louis, MO) for 30 minutes at 37 ?C. After 
coating, the excess fibronectin was removed, and the surfaces were rinsed with 37 ?C 
PBS. Cells were seeded at a density of 5?104 cells/cm2 and then cultured for 5-9 days. 
On days 2-10, cells were treated with medium containing cAMP supplements: 250 ?M 
of 8-CPT-cAMP (Abcam, ab120424) and 17.5 ?M of RO-20-1724 (0415, Tocris 
52 
 
 
Bioscience, Bristol, United Kingdom). Once supplemented medium was introduced to 
the cells, all of the following medium changes contained the supplements. 
4.2.3 Cell Viability and Metabolic Activity Assays, and Immunostaining 
Briefly, on day 1 of experiments, HBMECs were plated at 5?104 cells/cm2 
within fibronectin-coated glass-bottom plates. The following day, the cAMP 
supplemented medium was introduced to the system and cells were grown for 48 hours 
to confluency. On day 4, cells were incubated with 0.25 ?M benzoporphyrin derivative 
(VP, U.S. Pharmacopeia, Rockville, MD) in the supplemented medium for 24 hours. 
On day 5, prior to PDP, cells were washed twice with PBS and the fresh supplemented 
medium was added to each well. For PDP, cells were irradiated with 690 nm red light 
(0-5 J/cm2, 6 mW/cm2, bottom illumination). Light activation was performed using the 
ML6600 series laser system (690?5 nm; 1.5 W max output power; SMA fiber output 
with SMA905 connector, Modulight, Tampere, Finland). Cell metabolic activity and 
cell viability were determined via 3-(4,5-dimethylthiazol-2-yl)-2,5-
diphenyltetrazolium bromide (MTT) assay and LIVE/DEAD? staining, respectively, 
at 24 hours or 72 hours after PDP. Surviving cells for PDP treated samples were 
normalized to a no treatment control. Cell confluency is defined as the number of cells 
stained with Calcein AM per frame, normalized to the no treatment control. Cell 
morphology was assessed 90 minutes after PDP (see Junction analysis). 
4.2.4 Immunostaining of Junction Proteins 
At 90 minutes after PDP, HBMECs were rinsed with 37 ?C PBS and fixed with 
1% formaldehyde in PBS (Thermo Fisher Scientific) for 20 minutes. Prior to 
53 
 
 
permeabilizing the cell membrane for 5 minutes with 0.25% TritonX-100 (Sigma-
Aldrich) in PBS, cells were washed three times for 5 minutes each, with room 
temperature PBS. After permeabilization, the PBS wash steps were repeated and the 
samples were blocked for 1 hour at room temperature with 2% goat serum (Abcam, 
Cambridge, United Kingdom) in PBS. Cells were incubated with primary antibodies 
against ZO-1 (rabbit polyclonal, Thermo Fisher Scientific, 61-7300, 1:500) and VE-
cadherin (mouse monoclonal, Santa Cruz, sc-9989, 1:50) in 2% goat serum overnight 
at 4?C. The following day, cells were washed and blocked in 2% goat serum for 1 hour. 
Cells were incubated with secondary antibodies goat anti-rabbit Alexa Fluor 488 
(Abcam, ab150077, 1:100) or goat anti-mouse Alexa Fluor 568 (Thermo Fisher 
Scientific, A-11004, 1:100), and  Hoechst (Thermo Fisher Scientific, H3570, 1:2500 or 
4 ?g/mL), in PBS for 1 hour at room temperature. Cells were washed prior to imaging. 
Note that all steps were performed under gentle rocking.  
4.2.5 Microscopy 
For the LIVE/DEAD? assay, samples were imaged using the Lionheart 
(BioTek) at 20X. For cell-cell junction experiments (fixed-cell), samples were imaged 
using a 60X oil objective on an inverted microscope (IX83 Olympus microscope, 
Olympus cellSens Software). Images were captured using the red, green, and blue 
filters. Images shown within the manuscript were enhanced via ImageJ for better 
visualization of the cell-cell junction. 
54 
 
 
4.2.6 Junction Analysis 
Cell morphology and cell-cell junction organization were quantified using the 
JAnaP, available for download at https://github.com/StrokaLab/JAnaP. Briefly, the cell 
perimeter was traced via ?waypointing? for ZO-1 images (green fluorescent channel, 
A488) which were then projected onto the images of VE-cadherin (red fluorescent 
channel, A568), as previously described in [223]. To isolate ZO-1 and VE-cadherin, 
threshold values of 15 and 5 were applied, respectively. Further guidance on how to 
identify threshold values can be found in the supplement of [105] and in the JAnaP 
User-Guide available using the link above. Next, the cell morphological parameters, 
including area, solidity, and circularity, were calculated. Specifically, solidity equals 
the area of the cell divided by the area of the convex (Figure 4.1a). Circularity equals 
4? times the cell area divided by the cell perimeter squared (Figure 4.1b). Junction 
morphology was quantified by calculating the percent of the cell edge presenting 
continuous or perpendicular junction [223]. The specific phenotypes were classified 
based on the length of the junction that overlapped the cell edge (> 15 pixels for 
continuous junction) and the relative aspect ratio with respect to the cell edge (> 1.2 
pixels for perpendicular junction, otherwise punctate). These values served as constant 
parameters when using the JAnaP.  
55 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 4.1 Cell shape factors. Equations and schematics represent how (a) circularity and (b) 
solidity are calculated. Cells are stained for VE- cadherin (red). Green dashes represent 
waypoints that trace the cell perimeter (p), showing the area of a cell (Acell). White circles 
represent the area of the cell convex (Aconvex).   
4.2.7 Western Blot Analysis 
VE-cadherin expression was analyzed using a standard Western blot analysis 
protocol (Bio-Rad). Briefly, HBMECs were seeded at a density of 5?104 cells/cm2 
within FBN-coated 35 mm petri dishes and grown to confluency. On day 2, cAMP 
supplemented medium was added to the cells. On day 4, cells were incubated with 0.25 
?M VP in the supplemented medium for 24 hours. On day 5, prior to PDP, cells were 
washed twice with PBS and the fresh supplemented medium was added to wells. For 
PDP, cells were irradiated with 690 nm red light (0 J/cm2 or 1.2 J/cm2, 6 mW/cm2). 
Prior to collection, cells were washed thrice with cold PBS. Lysates were collected 90- 
and 24-hours post PDP by scrapping the cells off the petri dish within cold PBS. After 
centrifugation, cells were lysed with cold radioimmunoprecipitation assay (RIPA) 
buffer containing 1x protease and phosphatase inhibitor cocktail (Thermo Fisher 
56 
 
 
Scientific). The total protein from cell lysates was quantified using a Bichoninic acid 
protein assay (Thermo Fisher Scientific). Cell lysates (20 ?g) were separated on precast 
4-12% Bis-Tris protein gels (NuPAGE) and electrotransferred onto PVDF membrane 
(Thermo Fisher Scientific). Membranes were blocked either with 5% milk/TBST or 
5% bovine serum albumin/TBST solution, for VE-cadherin and GAPDH, respectively. 
Proteins were sequentially detected using an anti-VE-cadherin antibody (1:1000, Santa 
Cruz sc-9989) according to the manufacturer?s guidelines. GAPDH (1:500, Cell 
Signaling #2118) was used as the loading control. Visualization of protein bands was 
achieved using chemiluminescence (SuperSignal, Thermo Fisher Scientific). The 
membranes were imaged with the FluorChem E System (ProteinSimple, San Jose, CA). 
Quantitative analysis of protein expression was done using ImageJ software.  
 
4.2.8 Liposome Synthesis and Characterization 
Liposomes, fluorescently labeled with rhodamine, were synthesized via freeze-
thaw extrusion as described previously [19, 175]. Briefly, 
dipalmitoylphosphatidylcholine (DPPC), cholesterol, 
distearoylphosphatidylethanolamine-methoxy polyethylene glycol (DSPE?PEG) and 
1,2-dioleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (DOPG) (Avanti Polar Lipids, 
Alabaster, AL, USA) were codissolved with 1,2-dipalmitoyl-sn-glycero-3-
phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (Also known as 16:0 Liss 
Rhod PE) in chloroform. A thin film was created via rotary evaporator overnight. The 
film was rehydrated with 1 mL of deionized water. The lipid suspension was subjected 
to freeze-thaw cycles (4?45?C). The dispersions were extruded through polycarbonate 
57 
 
 
membranes (0.1 ?m pore size) at 42?C to form unilaminar vesicles. The final 
concentration of the rhodamine was checked using a fluorescence standard curve 
measured using a BioTek Synergy Neo2 plate reader (Excitation/Emission: 560/596 
nm). The Omni (Brookhaven, Holtsville, NY, USA) measured particle size of 
monodispersed rhodamine-liposomes to be ~100 nm in diameter (Table 4.1).  
4.2.9 Transwell Permeability Assay 
For permeability studies, HBMECs (5?104 cells/cm2) were seeded on transwell 
inserts that had been pre-coated with fibronectin. On the next day, cAMP supplemented 
medium was added, and cells were allowed to grow to confluency. On day 2 or 4, VP 
(0.25 ?M)-containing medium was added to the upper chamber for 24 hours of 
incubation. Following this, cells were washed twice with PBS, fresh medium was 
added, and light was delivered to initiate PDP (690 nm, 0.6 J/cm2 or 1.2 J/cm2, 6 
mW/cm2, bottom illumination). Immediately following PDP, 400 ?L of FITC-Dextran 
(1 mg/mL, 70 kDa, Sigma-Aldrich) or rhodamine-loaded liposomes (5 ?M) was added 
to the upper (apical) chamber and allowed to diffuse across the membrane into the 
lower (basolateral) chamber. At different time points, the medium in the lower chamber 
was collected and the fluorescence signal was measured using a BioTek Synergy Neo2 
plate reader (Excitation/Emission: 492/518 nm for FITC, or 560/596 nm for 
rhodamine). When evaluating the permeability of FITC-Dextran, on each of the 
following time points, the upper chamber was replenished with 400 ?L of fresh FITC-
dextran solution (1 mg/mL) and the lower chamber was also replenished with fresh 
medium (800 ?L), in order to continue daily permeability evaluation. When evaluating 
the permeability of rhodamine-loaded liposomes, 30 minutes prior to collection, the 
58 
 
 
upper chamber was replenished with 400 ?L of fresh rhodamine-loaded liposomes (5 
?M) and the lower chamber was replenished with fresh medium (800 ?L). The apparent 
permeability coefficients (Papp, cm/min) of FITC-dextran solution (1 mg/mL) or 
rhodamine-loaded liposomes (5 ?M) were from the following equation: 
Papp = Vr ? (dC / dt) / (C0?A) 
Vr is the volume of the basolateral chamber (cm
3). dQ/dt is the slope of the cumulative 
concentration of the FITC-dextran (or rhodamine-liposome) in the basolateral chamber 
over time (?mol/L/min). C0 is the initial concentration of FITC-dextran (or rhodamine-
liposome) in the apical chamber (?mol/L). A is the membrane surface area of the inserts 
(cm2).  
4.2.10 Statistical Analysis 
Data analysis was performed using PRISM GraphPad 8 [224]. Both the 
Anderson-Darling test and D'Agostino & Pearson test were used to determine the 
normality of the data sets. When examining cell and junction morphology, the Kruskal-
Wallis test followed by Dunn?s multiple comparison test were used to evaluate 
statistically significant difference. An alpha value of 0.01 was used as the cut-off value 
to determine significance, where ** p < 0.01, *** p < 0.001, and **** p < 0.0001. 
Three separate trials were conducted for all experiments. No batch effects were found. 
For transwell permeability experiments, a two-way ANOVA with multiple 
comparisons (mixed effects analysis) was used to assess statistically significant 
difference among groups. An alpha value of 0.05 was used as the cut-off value to 
determine significance, where * p < 0.05, ** p < 0.01, *** p < 0.001, and **** p < 
0.0001.  These analysis strategies were adopted from previously published works [108]. 
59 
 
 
 
4.3 Results & Discussion 
4.3.1 Sub-cytotoxic PDP in HBMECs 
For safe and effective drug delivery to the brain, it is of paramount importance 
to disrupt the BBB locally and transiently without causing endothelial cell death or 
vascular shutdown. Typically, PDT involves the high optical energy (e.g. 80?300 
J/cm2) activation of PS in order to produce sufficient 1O2 that can collapse the 
vasculature and induce cell apoptosis or necrosis [134, 140-143, 225]. Recently, we 
have shown that PDP, with slightly lower optical energy (75 J/cm2) and VP PS, can 
improve vasculature permeability to nanomedicine in pancreatic tumors without 
causing normal tissue damage [19, 175, 222]. Other groups have shown the ability to 
permeabilize the BBB within mice using lower optical energies (10-40 J/cm2) and the 
prodrug ALA, however, vascular damage and edema often occur [15-17]. In this study, 
we used LIVE/DEAD? assay and MTT (metabolic-based) assay to determine the 
threshold of PDP in HBMECs in vitro. To recapitulate the in vivo basement membrane 
and enhance the brain endothelial phenotype, HBMECs were plated on fibronectin and 
treated with 8-(4-chlorophenylthio) adenosine-3?,5?-cyclic monophosphate sodium salt 
(CPT-cAMP) and RO-20-1724. The basal lamina fibronectin protein was selected over 
7 other extracellular matrix components based on our prior work that showed 
fibronectin-coating results in the greatest HBMEC cell-cell junction expression [108, 
226]. cAMP and RO-20-1724 were introduced to enhance the junction phenotype, 
inhibit Rho/ROCK signaling in endothelial cells, and limit the formation of stress fibers 
by blocking myosin light chain phosphorylation [105, 108, 227].  
60 
 
 
Here, free-form VP was incubated with HBMECs at a concentration of 0.25 
?M for 24 hours prior to 690 nm light activation (0?5 J/cm2, 6 mW/cm2). At 24 hours 
post-PDP, the viability of HBMECs was assessed by LIVE/DEAD? assay. In a 
separate study, 72 hours post-PDP, mitochondrial activity was determined via MTT 
assay. Figure 4.2 shows that PDP at light doses of ? 2.5 J/cm2 had minimal effects on 
the viability and metabolic activity of HBMECs. At 5 J/cm2, PDP significantly 
decreased the mitochondrial activity by 35.7?6.4% (Figure 4.2a), induced 17.7?6% 
cell death (Figure 4.2b), and reduced the confluency of HBMECs by 63.6% (Figure 
4.2c, Figure 4.3). It is well established that VP localizes to the mitochondria and 
induces depolarization of mitochondrial membrane potential efficiently upon light 
illumination in cells [138, 139]. Therefore, it is not surprising that PDP-treated 
HBMECs experience a larger loss in mitochondrial activity (by MTT assay) than cell 
membrane integrity (by LIVE/DEAD?). Based on these results, optical energies of 
0.06, 0.3, 0.6, and 1.2 J/cm2 (at a fixed irradiance of 6 mW/cm2) were selected for the 
following experiments, while VP concentration is fixed at 0.25 ?M. 
61 
 
 
Figure 4.2 PDP is Sub-cytotoxic to HBMECs. HBMECs were incubation with VP (0.25 ?M) 
for 24 hours followed by light activation (690 nm, 0?5 J/cm2, 6 mW/cm2). The metabolic 
activity and viability of HBMECs were examined via (a) MTT assay (b) Live/Dead? imaging 
at 72 hours and 24 hours post-PDP, respectively. (c) Representative images from the 
Live/Dead? assay (green channel: Calcein AM, red channel: ethidium homodimer-1, EthD-1) 
show a confluent monolayer up to a PDP dose of 1.2 J/cm2 (scale bars = 100 ?m). Cell viability 
and metabolic activity were only significantly reduced by the 5.0 J/cm2 PDP dose. For 
Live/Dead? analysis, a One-Way ANOVA with multiple comparisons was used to calculate 
significant differences, where n.s. indicates not significant (p > 0.05), * p < 0.05. N=3, where 
N equals the number of trials performed. For MTT analysis, the Kruskal-Wallis test with a 
Dunn?s multiple comparison test was used to calculate significant differences, where n.s. 
indicates not significant (p > 0.05), * p < 0.05. N=6, where N equals the number of wells 
assessed over 3 trials. Box plots show the mean and the likely range of variation. Error bars 
show the maximum and minimum values.    
 
 
 
 
 
 
 
62 
 
 
 
 
 
 
 
 
Figure 4.3 Low dose PDP does not Impact Cell Confluency. HBMECs were incubated with 
VP (0.25 ?M) for 24 hours followed by light activation (690 nm, 0?5 J/cm2, 6 mW/cm2). The 
confluence of HBMECs was determined from Live/Dead? imaging at 24- post-PDP. The 
number of cells stained with Calcein AM (green channel) per frame was normalized to the 
control group. A One-Way ANOVA with multiple comparisons was used to calculate 
significant differences, where n.s. indicates not significant (p > 0.05), * p < 0.05, ** p < 0.01, 
*** p < 0.001, and **** p < 0.0001. N > 10, where N equals the number of images captured 
over 3 trials. Box plots show the mean and the likely range of variation. Error bars show the 
maximum and minimum values. 
4.3.2 PDP Slightly Alters Cell Morphology and F-actin Organization 
 
To better understand how PDP impacts the brain endothelium, we first 
examined the impact of PDP on HBMEC morphology (i.e., area, solidity, and 
circularity) and F-actin organization using the well-established JAnaP. We observed 
that PDP (0.06?1.2 J/cm2) moderately increases the circularity and solidity of 
HBMECs by 3?13%, suggesting cells were under stress (Figure 4.4a,b). However, this 
increase is modest compared to dying cells (cells treated with 5 J/cm2) that had on 
average a 23.5% increase in circularity (data not shown). PDP did not impact the 
HBMECs area (Figure 4.4c) nor perimeter (Figure 4.5). At the higher light doses of 
0.6 and 1.2 J/cm2, PDP-induced F-actin reorganization from cortical bands near the cell 
edge to stress fibers throughout the HBMECs (Figure 4.4d). It appeared VE-cadherin 
also showed a change in morphology, which motivated us to investigate further. Our 
new findings agree with previous observations by Ota et al. and others, which showed 
that the reactive oxygen species produced by PDT can induce morphological changes 
63 
 
 
and actin stress fiber formation in human umbilical vein endothelial cells [104, 228, 
229]. Further, these reports showed that actin stress fiber reorganization from cortical 
bands to stress fibers in human umbilical vein endothelial cells was also accompanied 
by the mislocalization of VE-cadherin [104]. In addition, stress fiber formation and 
induced permeability could be mitigated with the use of inhibitors. Forskolin/rolipram 
or 8CPT-2?O-Me-cAMP inhibit further reaction of phosphorylated RhoA, which 
reduced stress fiber formation and maintained the baseline permeability. 
 
 
 
 
 
64 
 
 
Figure 4.4 PDP Alters Cell Morphology and the Organization of F-actin. HBMECs were 
grown to confluency and primed with 690nm red light (0?1.2 J/cm2, 6 mW/cm2). At 90 minutes 
post priming, cells were fixed and stained for F-actin and VE-cadherin. (a-c) Cell shape factors. 
All PDP light doses modestly increase HBMEC (a) circularity and (b) solidity. (c) PDP had no 
impact on HBMEC area. (d) Representative fluorescence images of F-actin (green) 
organization. VE-cadherin (red) staining is used to show the cell edge. PDP light doses ? 0.6 
J/cm2 resulted in the rearrangement of F-actin from cortical bands near the cell boundary to 
stress fibers throughout the cell. All scale bars are 25 ?m. The Kruskal-Wallis test with a 
Dunn?s multiple comparison test was used to calculate significant differences, where n.s. 
indicates not significant (p > 0.05), ** p < 0.01, *** p < 0.001, and **** p < 0.0001. The 
significance between the experiment group and the control is presented. N > 105, where N is 
the number of cells, from 3 different trials. Box plots show the mean and the likely range of 
variation. Error bars show the maximum and minimum values. 
   
 
 
65 
 
 
 
 
 
 
Figure 4.5 PDP does not Alter Cell Perimeter. 90-minutes post priming (0-1.2 J/cm2, 6 
mW/cm2), cells were fixed and stained for VE-cadherin and ZO-1. The cell perimeter was traced 
via ?way pointing? within the JAnaP. The Kruskal-Wallis test with a Dunn?s multiple 
comparison test was used to calculate significant differences, where n.s. indicates not significant 
(p > 0.05). The significance between the experiment group and the control is presented. N > 
105, where N is the number of cells, from 3 different trials. Box plots show the mean and the 
likely range of variation. Error bars show the maximum and minimum values. 
4.3.3 PDP Alters the Endothelial Cell-cell Junction Morphology 
 
Among the 40 different transmembrane and scaffold proteins that form an 
endothelial cell-cell junction in the brain, VE-cadherin and ZO-1 are two of the 
important and foundational proteins that regulate BBB permeability. VE-cadherin is 
the primary transmembrane structural adhesive protein present in adherens junctions of 
HBMECs, prior to the formation of tight junctions, thus deeming it the junctional 
backbone [230-232]. ZO-1 is a scaffolding protein that regulates the recruitment and 
assembly of tight junction proteins (e.g., occludins and claudins) at the cytoplasmic 
surface of intercellular junctions [233]. It is also well established that rearrangement of 
VE-cadherin and ZO-1 regulates vascular permeability [234-238]. As cell-cell 
junctions mature, VE-cadherin and ZO-1 reorganize from a serrated perpendicular 
geometry to a continuous strand linear to the cell edge [107-109].  
 We next investigated whether PDP could reverse the maturity of endothelial 
cell-cell junction through the rearrangement of VE-cadherin and ZO-1. Similar to the 
previous experiments, HBMECs were seeded in FBN-coated wells and treated with 
66 
 
 
cAMP to start with mature cell-cell junctions [108]. Specifically, we examined the 
percentage of junction coverage per cell, as well as the pattern of junctions present at 
the cell edge (i.e., continuous mature junctions, perpendicular immature junctions, or 
punctate immature junctions) (Figure 4.6a). Representative fluorescence images 
(Figure 4.6b) show that PDP can reduce junctional coverage at the edge of the cell and 
significantly alter junction morphology. The changes in the HBMEC junction 
phenotype in response to PDP at various light energy densities (fluences) were further 
quantified using JAnaP as described by us previously (Figure 4.7). JAnaP analysis 
revealed that PDP modestly reduced the coverage of VE-cadherin (72.4?0.8%) and 
ZO-1 (64.6?0.7%) at the cell edge by 10-20% in a light dose-dependent manner 
(Figure 4.7a,b). This decrease in junctional coverage induced by PDP was 
accompanied by changes in junction morphology. We found that PDP not only 
decreases the mature VE-cadherin continuous junctions by ~17% (from 56.58?1.13% 
to 46.71?1.33%) (Figure 4.7c) but also increases the immature VE-cadherin 
perpendicular junctions by ~46% (from 7.91?0.26% to 11.61?0.38%) in a light dose-
dependent manner (Figure 4.7e). Similar trends were observed with ZO-1, where PDP 
decreases the mature ZO-1 continuous junctions by ~27% (Figure 4.7d) and increases 
the immature ZO-1 perpendicular junctions by ~38% at 1.2 J/cm2 (Figure 4.7). Further 
analysis revealed that PDP (0.3-1.2 J/cm2) increases the protrusion length (e.g. tip-to-
tip distance [223]) of ZO-1 perpendicular junction, but not VE-cadherin perpendicular 
junction (Figure 4.8). The changes in perpendicular junction protrusion could be a 
result of F-actin stress fibers pulling on the connected tight junction proteins (ZO-1), 
but not adherens junction proteins (VE-cadherin). It has been reported by others that 
67 
 
 
PDT could cause the internalization of VE-cadherin and intercellular gaps, resulting in 
the disappearance and discontinuous junctions at the cell edge [104]. Interestingly, PDP 
did not significantly alter the amount of punctate, or dotted, junctions for VE-cadherin 
and ZO-1 (p > 0.05) (Figure 4.7g,h). The number of punctate junctions was most likely 
unchanged after PDP, as cells were able to remain in constant contact with each other. 
Previous reports have shown that endothelial monolayer confluency impacts 
VE-cadherin expression and pattern [239-241]. Zhang et al. also showed that PDT 
downregulates the expression of VE-cadherin at the BBB in mice [17]. If PDP induced 
large gaps in the cell monolayer, as a result of cell death, it is speculated that 
downregulation of VE-cadherin would occur via clathrin-coated endosomal 
internalization [242]. Here, western blot analysis shows that PDP did not significantly 
alter the expression of VE-cadherin in HBMECs (Figure 4.9). The above results 
demonstrate that PDP not only induces cellular stresses as evident by increased stress 
fiber formation, roundness, and solidity, but also promotes morphological changes to 
junction proteins. Most importantly, PDP can shift a stable endothelial monolayer into 
an immature state while maintaining HBMEC viability. 
 
68 
 
 
 
Figure 4.6 PDP Modulates Endothelial Cell-cell Junction Phenotype. At 90 minutes post-
priming, cells were fixed, stained, and imaged. (a) Representative fluorescence image of PDP 
modulated HBMEC with stained nuclei (blue), ZO-1 (green), and VE-cadherin (red). White 
arrows show regions of different cell-cell junction phenotypes: continuous linear junctions, 
perpendicular junctions, and dotted punctate junctions. (b) Representative fluorescence images 
of non-treated cells (0 J/cm2) and photodynamic primed cells (0.06?1.2 J/cm2). PDP generates 
immature HBMEC monolayers by decreasing linear continuous junctions and increasing 
perpendicular junctions, in a light dose-dependent manner. All scale bars are 20 ?m. 
69 
 
 
Figure 4.7 Quantitative Analysis of Junction Phenotype. At 90 minutes post-priming (0?
1.2 J/cm2, 6 mW/cm2), cells were fixed, stained, and analyzed with the JAnaP. (a, b) The 
percent of the cell edge that is covered by a junction, for VE-cadherin and ZO-1, respectively. 
Edge presentation of continuous (c, d), perpendicular (e, f), and punctate (g, h) junctions for 
VE-cadherin and ZO-1, respectively. PDP resulted in a light dose-dependent decrease in mature 
junctions and an increase in unstable junctions. The Kruskal-Wallis test with a Dunn?s multiple 
comparison test was used to calculate significant differences, where n.s. indicates not 
significant (p > 0.05), ** p < 0.01, *** p < 0.001, and **** p < 0.0001. The significance 
between the experiment group and the control is presented. N > 105, where N is the number of 
cells assessed over 3 trials. Box plots show the mean and the likely range of variation. Error 
bars show the maximum and minimum values. 
 70 
 
 
Figure 4.9 PDP Impact on Protrusion Length. PDP did not alter the (a) protrusion length 
(i.e., tip-to-tip distance) of VE-cadherin perpendicular junctions, (b) but modulated the 
protrusion length of ZO-1 perpendicular junctions. At 90 minutes post-priming (0-1.2 J/cm2, 6 
mW/cm2), cells were fixed and stained. The JAnaP calculated the protrusion length of the 
perpendicular junctions after cells were way pointed. The Kruskal-Wallis test with a Dunn?s 
multiple comparison test was used to calculate significant differences, where n.s. indicates not 
significant (p > 0.05). The significance between the experiment group and the control is 
presented. N > 105, where N is the number of cells, from 3 different trials. Box plots show the 
mean and the likely range of variation. Error bars show the maximum and minimum values. 
 
Figure 4.8 PDP does not Alter Total Junction Protein Expression. HBMECs were grown 
to confluency, incubated with VP (0.25 ?M) for 24 hours, and primed (690 nm, 1.2 J/cm2, 6 
mW/cm2). Representative immunoblotting of VE-cadherin confirms protein expression was 
unchanged at 90 minutes and 24 hours post-priming. GAPDH was used as the loading control. 
N=4, where N is the number of gels ran over 2 lysates. 
 
 
4.3.4 PDP Induces HBMEC Permeability to FITC-dextran 
We have previously demonstrated that the number of discontinuous VE-
cadherin and ZO-1 junctions correlates positively with HBMEC permeability [108]. 
Similarly, Li et al. showed that a change in the pattern of ZO-1 (from ?linear? junction 
to a ?discontinuous, segmented and dotted? phenotype) is associated with increased 
71 
 
 
permeability in HBMECs [227]. Knowing that PDP can modulate the morphology of 
cell-cell junctions (Figures 4.6 and 4.7), we next examined the ability of PDP to 
improve HBMEC permeability to FITC-dextran or rhodamine-liposomes using a 
transwell model. At different time points after PDP, the amount of FITC-dextran (or 
rhodamine-liposome) that diffuses from the apical chamber into the basolateral 
chamber was quantified using a plate reader and the apparent permeability coefficient 
(Papp, cm/min) was calculated. 
Without PDP (0 J/cm2), a baseline diffusion of FITC-dextran into the 
basolateral chamber was observed (Figure 4.10a). The average Papp of FITC-dextran 
is ~7.5?10-5 cm/min, similar to that observed by others (Figure 4.10b) [243, 244]. By 
day 7, the Papp of FITC-dextran in the untreated HBMECs control group began to 
increase. This observation is consistent with other?s reports showing that prolonged 
activation of cAMP may cause a delay in the repression of Ras-related protein (R-Ras), 
resulting in the disruption of VE-cadherin and endothelial cell barriers [245]. PDP at 
0.6 J/cm2 and 1.2 J/cm2 significantly improved the diffusion of FITC-dextran across 
the HBMEC monolayer by ~2.6-fold and 4-fold, respectively, in a dose-dependent 
manner (Figure 4.10a). The Papp of FITC-dextran was found to be the highest at 2-3 
days post-PDP. On day 2, transwells primed with 1.2 J/cm2 had approximately 40.86% 
of their starting FITC-dextran diffuse into the basolateral chamber. By days 6 and 9, 
the Papp of FITC-dextran in the PDP groups (0.6 and 1.2 J/cm
2) were found to be similar 
to that of the control group (0 J/cm2). The transwell models that received the higher 
optical energy (1.2 J/cm2) took an additional 3 days to recover presumably due to 
increased cell contractility and number of immature junctions. Although this was an in 
72 
 
 
vitro model, our findings are consistent with previous findings showing that PDP-
assisted endothelial opening persists at least up to 3 days in vivo [175]. These results 
also suggest that PDP dose can be customized to control the degree and duration of 
endothelium permeabilization.  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 4.10 PDP Improves HBMEC Monolayer Permeability to FITC-dextran. (a) Cumulative 
FITC-dextran that traversed the HBMEC transwell model. Over 9 consecutive days, PDP (0.6-1.2 
J/cm2, 6 mW/cm2) resulted in 2.6?4-fold more FITC-dextran crossing into the bottom transwell 
chamber.  (b) Apparent permeability coefficient, Papp. PDP temporarily and reversibly induced a 
relatively high degree of permeability in HBMECs. By day 6 and 9, the integrity of the monolayers 
recovered. A two-way ANOVA with multiple comparisons was used to determine significance, where 
*P < 0.05, **P < 0.01, ***P < 0.001, n.s.: nonsignificant.  
 
We have previously shown that PDP can improve the permeability of ?leaky? 
tumor vasculature to liposomes by 11-fold in vivo [175]. We next tested if PDP also 
improved rhodamine-liposomes, ~100 nm in diameter (Table 4.1), penetration across 
the HBMEC monolayer cultures that have a particularly high degree of junctional 
tightness. The permeability of rhodamine-liposomes was monitored up ? to ~48 hours 
73 
 
 
(Figure 4.11), as the clinically relevant liposomes typically have a circulation half-life 
of 24?48 hours [246, 247]. Without PDP (0 J/cm2), the average Papp of rhodamine-
liposomes is ~1.1?10-7 cm/min. PDP at 0.6 J/cm2 significantly improved the Papp of 
rhodamine-liposomes by 14.6-, 9.8-, and 5.0-fold at 1, 25 and 49 hours, respectively, 
compared to the control (0 J/cm2). This improvement in the Papp of rhodamine-
liposomes was even more pronounced when using PDP and 1.2 J/cm2 (38.5-, 18.1-, and 
11.5-fold at 1, 25 and 49 hours, respectively). We observed that only less than 0.09% 
of the rhodamine-liposomes diffused into the basolateral chamber. This tightness 
properties displayed in our model was found similar to others in vitro model [248]. At 
1.2 J/cm2, PDP facilitated the leakage of up to 3.43% of the liposomes into the 
basolateral chamber over a 30-minute incubation period. This significant (30-fold) 
improvement in drug penetration across the endothelial barrier suggests that PDP is a 
promising modality for enhancing drug devilry across the BBB in vivo, where less than 
0.14%ID/g of the injected liposomes can cross the rat BBB [249-253]. While ALA-
mediated PDT and focused ultrasound [104, 254, 255] can also improve the drug 
delivery across the endothelial monolayer by 2-5.3 fold, these modalities are often 
cytotoxic and can potentially lead to undesired normal tissue destruction. In addition, 
liposomes can be designed to penetrate the BBB. For example, Dai et al. showed only 
0.58% of liposomes decorated with CDX peptide could cross a monolayer of rat 
primary brain capillary endothelial cells over a 3-hour period [248]. 
This is the first study to demonstrate that PDP can significantly improve 
endothelial permeability to both macromolecules and nanoparticles in a transwell 
model. The uniqueness of PDP lies in its spatiotemporal selectivity, which offers the 
74 
 
 
opportunity to confine endothelial permeabilization to the site where the light is 
directed. This spatiotemporal control of PDP may further reduce normal brain tissue 
damages and warrants further investigation in animals. While it is true that light 
penetration depth (i.e., the depth at which the incident optical energy drops to 37%, 
1/e) is typically 1-4 mm in the brain, we have recently shown that diffused red light 
(i.e., the remaining 37% optical energy) can reach up to 1.5-2 cm deep in rat brains to 
activate VP for PDP [22]. This suggests that intraoperative PDP is sufficient to manage 
brain diseases that are 1-2 cm in depth (e.g., the unresectable, invasive margin of most 
brain cancers). For deeper brain tissues, light can be delivered to the entire target using 
stereotactically placed fibers in the clinic (similar to laser probes currently used for 
interstitial thermotherapy) [80, 256-259]. In this study, the clinically relevant, longer 
wavelength PS, VP, (690 nm, deeper light penetration) was selected for PDP. This is 
an alternative to ALA (Gliolan?)-induced protoporphyrin IX (PpIX, 635 nm), which 
is currently approved for fluorescence-guided surgery of brain tumors [149, 260] and 
is generally reserved for treating topical lesions [140, 261]. In fact, we have shown that 
690 nm light activation of VP induces PDP effects at further depths in rat brain (1.5-2 
cm), compared to 635 nm light activation of PpIX (? 1 cm) [22]. Therefore, an 
immediate future direction would be to compare the efficacy of PDP for BBB opening 
using both FDA-approved VP and Gliolan? in vivo. 
 
Table 4.1 Physical Parameters of Rhodamine-liposomes. 
 
75 
 
 
 
Figure 4.11  PDP Improves HBMEC Monolayer Permeability to Rhodamine-liposomes. Priming 
(0.6-1.2 J/cm2, 6 mW/cm2) the transwells resulted in the apparent permeability coefficient, Papp, of the 
HBMECs to increase by as much as 38.5-fold, allowing rhodamine-nanoliposomes to diffuse into the 
basolateral chamber. A two-way ANOVA with multiple comparisons was used to determine 
significance, where n.s. indicates not significant (p > 0.05), * p < 0.05, ** p < 0.01, *** p < 0.001, and 
**** p < 0.0001. The significance between the experiment group and the control is presented. N ? 5, 
where N is the number of inserts measured over 3 trials. Error bars show SEM. 
4.4 Conclusion 
5. In summary, the complexities of the brain necessitate the innovation of a drug 
delivery strategy capable of overcoming the neurovascular unit while leaving 
normal brain cells undamaged. An emerging paradigm in brain drug delivery 
suggests that stimuli-responsive strategies can more accurately control BBB 
permeability at the right time, right place, and for the appropriate duration. Both 
PDT and focused ultrasound hold great clinical promise for overcoming the BBB. 
However, their cytotoxic nature and associated adverse events limit the clinical 
application and efficacy of drug delivery. To address these challenges, we applied 
a sub-lethal, photodynamic modality to reverse the mature brain endothelial cell?
cell junction. The photodynamic priming approach is inspired by recent advances 
in tumor vascular permeabilization, which improves drug delivery to pancreatic 
tumors. Here, for the first time, we showed that PDP could significantly alter the 
76 
 
 
endothelial cell?cell junction phenotype (i.e., continuous, punctate, or 
perpendicular), and improve nanoparticle penetration across the tight endothelium 
by up to 30-fold. In Chapter 7, we will apply PDP to open the BBB within healthy 
rats. Future work will involve further characterization of multi-cycle PDP in cells 
and in animal models and will address the benefit of using a longer wavelength 
photosensitizer. In a small pilot study, we examined the colocalization of CD34 and 
claudin 5 after PDT treatment in healthy rat brain (PDT parameters discussed in 
Chapter 7). Compared to the unaffected left-brain hemisphere which had a 
colocalization of 96.2%, PDT decreased colocalization to 78.4%. The impact of 
PDP on other components of the neurovascular unit (e.g., pericytes, neurons), as 
well as the PS-light interval, will also be the subject of future studies. Building on 
our successes using PDP for pancreatic cancer treatment, this study opens up new 
avenues for developing brain targeting drug delivery systems. 
77 
 
 
6. Chapter 5. The Development of Verteporfin 
Biomolecules for Anti-glioma Photodynamic 
Therapy3 
5.1 Introduction 
As mentioned in Sections 2.3.2 and 3.1, free-form VP cannot be used clinically for 
treatment. VP is hydrophobic and in aqueous solutions it will form large uncontrollable 
agglomerations that will negatively impact pharmacokinetics and 1O2 production. VP 
requires a carrier system for clinical delivery. 
Light-activatable biohybrid conjugates offer great promise for anti-cancer PDT 
due to their ability to improve pharmacokinetics, minimize clearance of 
photosensitizing agents, and enhance tumor-specific cytotoxic effects [262]. PDT is a 
light-based treatment modality that is mechanistically distinct from conventional 
chemotherapy, radiation, and thermal ablation [126, 263]. Cytotoxicity related to PDT 
is governed by the localization of the photosensitizer, spatial confinement of near-
infrared light activation, and the production of ROS (e.g., 1O2, H2O2, O
?-
2 , ?OH) that 
confer toxicity to cellular and subcellular targets [126, 263]. The direct damage to 
subcellular organelles (e.g., mitochondria, endoplasmic reticulum, lysosome) allows a 
more channeled activation of programmed cell death pathways (e.g., apoptosis, 
necroptosis, autophagy), and therefore bypasses many other cell-death signaling 
pathways required for chemotherapy and radiation to be effective [126, 263]. As a 
                                                 
3 This chapter was adapted from Inglut C.T.#, Baglo Y.#, Liang B.J., et al., ?Systematic Evaluation of 
Light-Activatable Biohybrids for Anti-Glioma Photodynamic Therapy?. J Clin Med. 2019 Sep; 8(9): 
1269. No special permission is required to reuse this article published by MDPI under the terms and 
conditions of the Creative Commons Attribution (CC BY) license 
(http://creativecommons.org/licenses/by/4.0/). Y.B. was a co-first author (# denotes equal contribution) 
on this article. She helped perform and analyze the experiments for Figures 5.4, 5.5a-c, 5.6, 5.8a, 5.9a, 
and 5.10c. B.J.L. aided in synthesis and characterization (Figure 5.1-5.3). 
78 
 
 
result, PDT has been shown to reverse chemoresistance, synergize with 
chemotherapeutics and biological agents, and overcome compensatory survival 
pathways used by numerous cancers to evade standard treatments [19, 175, 197, 264-
266]. 
In the past few decades, considerable attention has been given to improving the 
biocompatibility of organic photosensitizing agents, including porphyrins, chlorins, 
and synthetic dyes [127, 128]. It is increasingly evident that photosensitizing 
biomolecules (PSBMs), based on chemical conjugation or association of 
photosensitizers with biomolecules, strongly influence the performance of a given 
photosensitizer in biological environments [262]. To harness the full potential of 
PSBMs for cancer treatment, it is of paramount importance to achieve a general and 
comprehensive understanding of how different types of biomolecules alter the uptake, 
subcellular localization, and the cytotoxicity of photosensitizers. However, the 
numerous studies examining different PSBMs are not readily comparable as they 
typically cover a vast array of photosensitizers and treatment conditions [262]. This 
study systematically examines the impact of a panel of biomolecules on the behavior 
of VP, a FDA-approved photosensitizer now being routinely tested for cancer treatment 
and imaging [145, 191]. 
Amongst the PSBMs that are currently available in the clinic, phospholipid-, 
polymer-, and antibody-based formulations are popular ways to improve the water 
solubility, pharmacokinetics, and biodistribution of photosensitizers; however, little is 
known about how these re-formulation strategies alter cellular uptake, subcellular 
localization, and the end effects in PDT regimes against GBM. In 2000, phospholipid-
79 
 
 
associated VP (i.e., Visudyne?) was approved by the FDA for PDT of wet age-related 
macular degeneration [145, 191]. Accordingly, Visudyne? is now being tested in 
several clinical trials evaluating PDT for pancreatic ductal adenocarcinoma 
(NCT03033225) [168], recurrent and metastatic prostate cancer (NCT03067051, 
NCT02939274), breast neoplasms (NCT02872064), and pleural malignancies 
(NCT02702700). In an early clinical study, Bown et al. tested the use of intravenous 
polyethylene glycol (PEG)-solubilized meso-tetrahydroxyphenyl chlorin for enhanced 
PDT against locally advanced, unresectable pancreatic cancer [267]. In this study, light 
was delivered to the tumor percutaneously under computerized tomographic guidance, 
which achieved a 100% response rate and a median overall survival of 9.5 months. 
Later on, Moore et al. employed Cremophor?-emulsified palladium-
bacteriopheophorbide (TOOKAD?) for successful vascular-targeted PDT against 
prostate cancer [268]. Regarding antibody-based PSBMs, Schmidt et al. were the first 
to conduct a clinical study using antibody-directed phthalocyanine conjugates for 
targeted PDT in patients with advanced ovarian carcinoma [269]. More recently, a 
PSBM-based system using a cetuximab-IR700 conjugate (RM-1929) was designed for 
photoimmunotherapy of head and neck cancer and showed encouraging results in a 
clinical trial (NCT02422979) [270]. While the primary motivation of re-formulated 
photosensitizers with PSBMs is to improve the photo- and physio-chemical properties 
of the agents in vivo, it is apparent that the complex interactions between cellular 
uptake, subcellular localization, and molecular targets are more likely to govern the 
observed phototoxicity of a given PSBM. 
80 
 
 
In this study, we prepared and characterized a series of PSBMs by conjugating VP 
to (i) 1-stearoyl-2-hydroxy-sn-glycero-3-phosphocholine (16:0 Lyso PC) 
phospholipid, (ii) PEG-distearoylphosphoethanolamine-polyethylene glycol (DSPE-
PEG) phospholipid-polymer conjugate, or (iii) anti-epidermal growth factor receptor 
monoclonal antibody cetuximab. The study of lipid-conjugated tetraphenylporphyrin 
photosensitizers was first reported by Komatsu et al. in 2002 to form spherical 
unilamellar vesicles with a diameter of 100 nanometers (nm) [271]. This work was later 
followed by studies from Zheng, Lovell, and others using different linking strategies, 
photosensitizers, and lipids to synthesize porphysome nanovesicles for effective light-
based therapy of tumors [196, 203, 272]. Studies from Soukous and Hamblin showed 
that the surface charge and PEGylation of Chlorin e6 conjugates alter tissue-localizing 
prosperities and play an important role related to the efficacy of PDT for ovarian cancer 
[273, 274]. Hasan and others pioneered covalent conjugation of VP to Cetuximab for 
targeted photoimmunotherapy and fluorescence imaging [275-277]. The highly self-
quenched VP conjugated to Cetuximab can be internalized by cancer cells through 
clathrin-mediated endocytosis and de-quenched via lysosomal proteolysis of the 
antibody [276, 278]. Here, using 16:0 Lyso PC-VP, DSPE-PEG-VP, Cetuximab-VP, 
and free-form VP, we present a full physical, photochemical, cellular uptake, and 
subcellular localization investigation under controlled experimental conditions. This 
chapter will explore how these properties relate to PSBM cytotoxicity in human GBM 
cells.  
81 
 
 
5.2 Methods 
5.2.1 PSBM Preparation and Purification 
Three types of PSBMs were synthesized by covalently linking VP to (i) 16:0 Lyso 
PC (MW. 532.68; Avanti-Polar Lipids), (ii) DSPE-PEG (MW. 2790.48; Avanti-Polar 
Lipids), and (iii) Cetuximab (MW. ~146k; Bristol-Myers Squibb). The 16:0 Lyso PC-
VP conjugate was prepared by crosslinking the carboxylic acid group of VP to the 
hydroxyl functional group of 16:0 Lyso PC via the esterification reaction as described 
by others [167] and us previously [196]. Briefly, 16:0 Lyso PC, VP, 1-ethyl-3-(3-
dimethylaminopropyl) carbodiimide, 4-(dimethylamino) pyridine, and N,N-
diisopropylethylamine were mixed in dichloromethane at a fixed molar ratio of 
1:5:50:25:60 (113 mg total) for 24 hours at room temperature. Dichloromethane was 
removed via rotary evaporation, and the residue was suspended in 5 mL of methanol 
prior to Sephadex? LH-20 gel chromatography column purification. Following which, 
methanol was removed via rotary evaporation and the purified 16:0 Lyso PC-VP was 
stored at -20?C. The DSPE-PEG-VP conjugates were prepared using carbodiimide 
crosslinker chemistry as described by others previously [279]. Briefly, 1-ethyl-3-(3-
dimethylaminopropyl) carbodiimide, VP, and DSPE-PEG were mixed chloroform at a 
molar ratio of 50:10:1 (20 mg total) for 24 hours at room temperature. Chloroform was 
removed via rotary evaporation, and the resulting thin film was hydrated with 1.5 mL 
of methanol for purification with a Sephadex? LH-20 gel chromatography column. 
The purified DSPE-PEG-VP conjugates were dried via rotary evaporation and then 
stored at -20 ?C storage. Conjugation of VP to Cetuximab was achieved using 
carbodiimide crosslinker chemistry as described by others and us previously [275, 278]. 
82 
 
 
Briefly, PEGylation of Cetuximab was carried out by adding mPEG-NHS in DMSO 
(200 ?L; 40,000 MW; 4 mg/mL) dropwise to Cetuximab (2 mL; 2 mg/mL) and allowed 
to stir overnight at 400 rpm at room temperature. PEGylated Cetuximab was reacted 
with the N-hydroxysuccinimidyl ester of VP (VP-NHS) at 1:9 molar ratios for 20 hours. 
The resulting Cetuximab-VP conjugates were purified using a Zebra spin desalting 
column (7kDa MWCO; Thermo) pre-equilibrated with 30% DMSO in PBS, and then 
washed in a 20 mL centrifugal filter tube (30kDa MWCO; Amicon) three times with 
5% DMSO in PBS prior to storage at 4?C. The purified 16:0 Lyso PC-VP and DSPE-
PEG-VP conjugates were analyzed using matrix-assisted laser desorption ionization-
time of flight mass spectrometry (MALDI-TOF MS; Bruker). The VP concentration of 
PSBM was estimated by absorbance in DMSO using the established molar extinction 
coefficient of VP in DMSO (~34,895 M-1 cm-1 at 687 nm) [19]. The Cetuximab-VP 
purity was confirmed by gel fluorescence imaging analysis of free VP following 
sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). The protein 
content of Cetuximab-VP was evaluated by bichoninic acid protein assay (Thermo). 
VP conjugation efficacy was determined by dividing the amount of antibody-
conjugated VP in the purified Cetuximab-VP by the total amount of VP added initially. 
5.2.2 Photophysical and Photochemical Characterizations 
The fluorescence-based optical characterization (Excitation/Emission: 422/650-750 nm) 
of PSBMs in PBS or DMSO was determined using a multi-mode microplate reader (BioTek). 
Fluorescence photoactivity is defined as the maximal fluoresce intensity of PSBM in PBS 
divided by the maximal fluoresce intensity of PSBM in DMSO. The singlet oxygen yield of 
PSBM was studied in 96-well plates as described previously [19, 278]. Briefly, VP and SOSG 
83 
 
 
were fixed at 5 ?M, and light (690 nm, 10 J/cm2, 10 mW/cm2) was delivered vertically. The 
microplate reader acquired the fluorescence signals of SOSG (Excitation/Emission: 504/525 
nm).   
5.2.3 Cell Culture 
Human GBM cell lines U251 and U87 were obtained from American Type Culture 
Collection (ATCC) and cultured in Eagle's Minimum Essential Medium cell culture 
medium (Cellgro), supplemented with 10% (v/v) FBS (Gibco), 100 U/mL penicillin 
and 100 ?g/mL streptomycin (Lonza), according to the manufacturer?s instructions. 
The GBM39 patient-derived xenograft was originally obtained from patients at the 
Mayo Clinic by Dr. Jann Sarkaria.  Molecular genetic alterations and patient tumor 
information for GBM39 has been previously determined and described [280]. GBM39 
was derived from the primary GBM tumor of a 51-year-old male prior to radiation, 
chemotherapy (TMZ), craniotomy, and Carmustine (BCNU) wafers (Gliadel). GBM39 
cells were cultured in DMEM/F12 medium using StemPro NSC SFM Kit 
(ThermoFisher A1050901) including neural supplement, bFGF, and EGF, 
supplemented with 100 U/mL penicillin and 100 ?g/mL streptomycin (Lonza), and 
5mM GltaMAX (GIBCO). GBM39 cells were cultured to 70% confluency in a Poly-
L-lysine (Sigma) and laminin (GIBCO) coated culture flask before seeding. All cell 
lines were maintained in 5% CO2 at 37?C. Cultures were tested mycoplasma free after 
resuscitation.  
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5.2.4 Photodynamic Therapy of Glioblastoma Cells Using PSBMs 
For PDT studies, U251 and U87 cells were plated in 35-mm Petri dishes at a cell 
density of 7.5?104 and 2.5?105 cells per dish, respectively, to allow overnight culture. 
Attached cells with 80-90% confluency were incubated in serum-free medium 
containing free VP or PSBMs at desired concentrations for 24 hours, at which time 
cells were irradiated with 690 nm red light (20 J/cm2, 150 mW/cm2, bottom 
illumination) in the fresh complete culture medium. Light activation was performed 
using the ML6600 series laser system (690?5 nm; 1.5 W max output power; SMA fiber 
output with SMA905 connector, Modulight). Cell viability was determined by MTT 
assay at 24 hours after 690 nm light irradiation. Cell survival fractions for PDT treated 
samples were normalized to the no treatment controls. The no treatment controls were 
incubated in serum-free medium with no exposure to free VP or PSBM?s and were not 
subject to irradiation. Alternatively, cells were collected 4 hours post-treatment and 
incubated with Annexin V FITC and propidium iodide (Abcam, ab14085) for flow 
cytometry analysis (BD FACSCelesta). Mitochondrial membrane potential (??m) was 
examined via tetramethylrhodamine ethyl ester (TMRE) assay (Abcam). Briefly, U251 
cells were cultured overnight in 96-well black wall plates at a density of 3,000 
cells/well, and then incubated with PSBM for 24 hours (0.15 ?M of free VP, 0.25 ?M 
of 16:0 Lyso PC-VP, 0.5 ?M of DSPE-PEG-VP, and 0.5 ?M of Cetuximab-VP). After 
incubation, cells were washed twice with PBS and exposed to 690 nm light (0 or 1 
J/cm2, 10 mW/cm2). After PDT, cells were incubated with TMRE (250 nM)-containing 
medium for 25 minutes, washed twice with 0.2% bovine serum albumin within PBS, 
and subjected to fluorescence readings (Excitation/Emission: 549/575 nm) using a 
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microplate reader (BioTek). Controls include no light irradiation and 150 ?M of 
carbonilcyanide p-triflouromethoxyphenylhydrazone (FCCP). Lysosome damage was 
examined via LysoTracker Red incorporation. Briefly, cells were incubated with 0.15 
?M free VP, 0.25 ?M 16:0 Lyso PC-VP, 0.5 ?M DSPE-PEG-VP, or 0.5 ?M 
Cetuximab-VP for 24 hours and irradiated with 690 nm red light (20 J/cm2, 150 
mW/cm2, bottom illumination). At 30 minutes after light treatment, cells were 
incubated with 100 nM LysoTracker Red in serum free medium for 15 minutes at 37?C 
and washed twice with PBS. At least 20,000 events were collected by the flow 
cytometer (BD FACSCelesta) using the BD Horizon PE-CF594 filter 
(Excitation/Emission: 561/610) and analyzed with FlowJo software.  
5.2.5 PSBM Uptake and Localization Studies in Glioblastoma Cells 
To evaluate PSBM uptake, U251 (or U87) cells were plated in 35-mm Petri dishes 
at 2?105 and 2.5?105 cells per dish, respectively. Attached cells with 80-90% 
confluency were incubated in culture medium containing free VP or PSBMs (0-1 ?M, 
1 mL) for 24 hours. Cells were washed twice with PBS, harvested with trypsin, and 
lysed in Solvable solution (0.5 mL) at 4?C. VP fluorescence intensity 
(Excitation/Emission: 435/690?20 nm) was measured using a microplate reader 
(BioTek) converted to moles of VP based on appropriate standard curves. Protein 
concentration was determined using the Pierce bichoninic acid protein assay. To verify 
PSBM uptake using fluorescence microscopy imaging (Lionheart, BioTek), cells were 
plated at the same density within a 24-well glass bottom plate, and incubated with 0.25 
?M free VP or PSBM for 24 hours. Before imaging, cells were washed twice with PBS, 
and nuclei were stained with Hoechst nucleic acid stain. Live cell imaging was 
86 
 
 
performed to confirm aldehydes did not impact analysis. Images were captured in 
quadruplet at specified locations for each well. Fluorescent intensity of VP was 
quantified and normalized. Confocal fluorescence imaging studies were performed to 
determine the subcellular localization of PSBMs in GBM cells. Briefly, cells were 
cultured overnight in 35-mm glass bottom dishes at 75,000 cells per dish and incubated 
with free VP or PSBMs for 24 hours before confocal imaging. After PBS washing, cells 
were stained with 1 mL 100 nM MitoTracker Green (M7514; Invitrogen) or 100 nM 
LysoTracker Red probes (DND-99; Invitrogen) for 30 min at 37 ?C. Hoechst was 
utilized to stain the nuclei of the cells prior to confocal fluorescence imaging (Olympus 
FluoView-1000 confocal microscope) using a 10X or 40X objectives. Excitation of 
Hoechst/VP, MitoTracker, and LysoTracker were carried out using 405 nm, 488 nm 
and 559 nm lasers, respectively, with appropriate filters (Hoechst: 430-470 nm; VP: 
650-750 nm; MitoTracker: 500-540 nm; LysoTracker: 570-620 nm). Profile plots were 
created using ImageJ software showing co-localization of PSBM and mitochondria or 
lysosomes [281]. 
5.2.6 Immunoblotting 
U251 cells (7.5?105 cell per 35-mm dish) were treated with PSBM for 24 hours 
(0.15 ?M of free VP, 0.25 ?M of 16:0 Lyso PC-VP, 0.5 ?M of DSPE-PEG-VP, and 
0.5 ?M of Cetuximab-VP) followed by irradiated at 690 nm (1 J/cm2, 10 mW/cm2, 
bottom illumination) in the fresh culture medium. At 6 and 24 hours post-PDT, cell 
lysates were collected in RIPA lysis buffer supplemented with 1% phosphatase and 
protease inhibitor cocktail. Protein lysates (20 ?g) of each sample were separated on a 
NuPAGE 4?12% precast Bis-Tris gel (Mini-tank system, ThermoFisher) and 
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transferred onto a polyvinylidene difluoride membrane. Subsequent to blocking with 
5% bovine serum albumin in TBST, the blot was incubated with primary antibody 
against LC3I/II (Cell Signaling, #4108) overnight at 4 ?C followed by incubation with 
its secondary antibody for 2 hours at room temperature based on the supplier?s advice. 
Visualization of protein bands was developed by chemiluminescence (SuperSignal, 
ThermoFisher) with exposure to Gel Imager (ProteinSimple). ?-actin serviced as 
loading control. 
5.2.7 Statistical Analyses 
Results are mean ? standard error of the mean (SEM). Statistical tests were carried 
out using GraphPad Prism. All experiments were carried out at least in triplicate. 
Specific tests and number of repeats are indicated in the figure captions. Reported P 
values are two-tailed. One-way ANOVA statistical tests and appropriate post hoc tests 
were carried out to avoid type one error.  
5.3 Results and Discussion 
5.3.1 Synthesis and Characterization of PSBMs 
16:0 Lyso PC-VP PSBMs were synthesized by an esterification reaction between 
the carboxyl groups of VP photosensitizers and the alcohol groups of 16:0 Lyso PC 
lipids. The hydrophobic VP placed along the alkyl side chain of 16:0 Lyso PC 
maintains an overall amphipathic structure of the 16:0 Lyso PC-VP (Figure 5.1a). To 
prepare DSPE-PEG-VP, VP was coupled to the polar PEG-modified head group of 
DSPE-PEG via amide coupling, ultimately changing the amphipathic structure by 
capping the hydrophilic PEG head with a hydrophobic VP molecule (Figure 5.1b). 
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MALDI-TOF mass spectrometer analysis of the purified 16:0 Lyso PC-VP conjugates 
showed a single primary product with a mass of 1196.4, compared to multiple products 
for the crude reaction mixture (Figure 5.2a). The observed peak mass for the purified 
DSPE-PEG-VP PSBMs was 3602.1 (Figure 5.2b), which is similar to others finding 
[279]. The final yields of 16:0 Lyso PC-VP and DSPE-PEG-VP were approximately 
20.1?3.8% and 13.0?3.7%, respectively. It is well established that 16:0 Lyso PC-VP 
and DSPE-PEG-VP can self-assemble in aqueous medium into a variety of nano- to 
micron-scale vesicles when using the extrusion method [196, 203, 279]. The structure 
of the vesicles formed is dictated by the concentration of PSBMs as well as the self-
assembling environment [196, 203, 279]. Here, our goal is to examine the cellular 
interaction of the monomeric PSBMs instead of their nano/micro-formulations. Under 
our experimental conditions, no nano- or micro-vesicles of 16:0 Lyso PC-VP or DSPE-
PEG-VP were detected when using dynamic light scattering technique. 
To prepare Cetuximab-VP conjugates (Figure 5.1c), 7 VP photosensitizers were 
attached to the lysine groups of each anti-EGFR antibody Cetuximab at a conjugation 
efficiency of 66.1?3.6%. PEGylation of Cetuximab before VP conjugation was 
necessary to prevent the aggregation of Cetuximab-VP conjugates [278]. We have 
previously confirmed that conjugation of 6-7 VP molecules per Cetuximab does not 
impair the selective binding and internalization of Cetuximab in EGFR positive cancer 
cells [278]. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis was used to 
identify and monitor the purity of Cetuximab-VP, which is less than 1% free VP. 
Similar to 16:0 Lyso PC-VP and DSPE-PEG-VP, Cetuximab-VP can be in 
89 
 
 
incorporated into nanoconstructs by linking them to the outer layer of nanoparticles to 
achieve targeting functionality [128, 278, 282]. 
 
 
Figure 6.1 Synthesis Reaction and Chemical Structure of PSBMs. (a) 1-stearoyl-
2-hydroxy-sn-glycero-3-phosphocholine (16:0 Lyso PC) was conjugated to VP via 
esterification reaction, resulting 1:1 ratio of 16:0 Lyso PC to VP. Carbodiimide 
crosslinker chemistry was employed to conjugate VP onto (b) PEG-
distearoylphosphoethanolamine-polyethylene glycol (DSPE-PEG) and (c) 
monoclonal antibody cetuximab (Cet). Final conjugates contain 1:1 and 1:7 ratio of 
DSPE-PEG-to-VP and Cetuximab-to- VP, respectively. Green circles are to represent 
VP photosensitizers.  
 
90 
 
 
Figure 6.2 MALDI-TOF mass spectrometer analysis of 16:0 Lyso PC-VP and DSPE-
PEG-VP. (a) Purified 16:0 Lyso PC-VP showed a single primary product with mass of 1196.4 
(red line) compared the multiple products for the crude reaction mixture (blue line) (yield: 
20.1?3.8%). (b) Purified DSPE-PEG-VP demonstrated a shift peak mass at 3602.1 (red line) 
compared to the crude reaction mixture (blue line) (yield: 13.0?3.7%, N?3) 
 
5.3.2 Photoactivity of PSBMs 
Conjugation of VP to biomolecules is an expedient strategy to improve the 
photoactivity of VP under biologically relevant conditions. In clinical PDT practice, 
systemic VP photosensitizers are commonly activated by 690 nm light because near-
infrared light penetrates deeper into the tissues than light of shorter wavelength [168]. 
Here, we showed that conjugation of VP to 16:0 Lyso PC, DSPE-PEG, or Cetuximab 
does not alter the Q band (690 nm) or the Soret peak (435 nm) of VP (Figure 5.3a). 
Red shift of the Q band was not anticipated, as the lipid conjugation reaction does not 
reduce the number of double bonds in the pyrrole rings of VP [283]. We further 
evaluated the fluorescence emission spectra (650-750 nm) of free VP and PSBMs (i.e., 
16:0 Lyso PC-VP, DSPE-PEG-VP, or Cetuximab-VP) in PBS and in dimethyl 
sulfoxide (DMSO), using an excitation wavelength of 435 nm. The aggregation of free 
VP in PBS led to a nearly complete (>95%) fluorescence quenching as compared to 
fully solubilized VP in DMSO (Figure 5.3b). DSPE-PEG-VP restored the fluorescence 
emission signal of VP molecules in PBS and was the most effective at retaining up to 
91 
 
 
15% photoactivity of the VP (Figure 5.3c). In contrast, 16:0 Lyso PC-VP demonstrates 
less retention of photoactivity compared to that of DSPE-PEG-VP. This is likely caused 
by the improved stability due to the presence of PEG in DSPE. Cetuximab-VP shows 
the least retention of photoactivity at ~5%. This can be explained by our previous report 
and others that Cetuximab-VP conjugates at 1:7 ratio display up to 7-fold VP static 
quenching [275, 276]. More importantly, these highly self-quenched VP molecules on 
Cetuximab can be de-quenched by cancer cells upon lysosomal proteolysis of the 
Cetuximab antibody. This unique ?tumor-activation? function of Cetuximab-VP can be 
used to improve the selective delivery of VP for a safer photoimmunotherapy as shown 
by others and us [276, 278]. 
Type II photosensitization is the dominant process in VP-based PDT and produces 
highly reactive singlet oxygen (1O2), which confers toxicity to nearby targets by 
oxidizing proteins and lipids [23]. In this study, 1O2 generation from light-activated 
PSBMs or free VP is monitored indirectly by using the fluorescence probe SOSG. We 
have previously confirmed that the VP fluorescence is negligible at the wavelength 
used to monitor the SOSG fluorescence [19]. Upon 690 nm light activation, the SOSG 
fluorescence intensity generated by DSPE-PEG-VP was at least 4 folds higher than that 
generated by free VP, Cetuximab-VP, and 16:0 Lyso PC-VP (Figure 5.3d). This 
correlates with our finding of DSPE-PEG-VP exhibiting the highest retention of 
photoactivity (Figure 5.3c). The SOSG fluorescence intensity generated from 
Cetuximab-VP was not significantly higher than that produced by free VP and 16:0 
Lyso PC-VP upon light activation (P > 0.05, Figure 5.3d). This is likely due to the 
fluorescence quenching shown in Figure 5.3c. This data suggests that DSPE-PEG-VP, 
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with relatively stronger fluorescence emission spectra, improves the 1O2 production 
under physiologically relevant conditions during light activation, compared to free VP, 
Cetuximab-VP, and 16:0 Lyso PC-VP; which may lead to a higher phototoxicity 
response for PDT. 
Figure 6.3 Photophysical and Photochemical Characterization of PSBMs. (a) 
Representative absorbance spectra of free VP, 16:0 Lyso PC-VP, DSPE-PEG-VP, and 
Cetuximab-VP in DMSO. Conjugation of VP on biomolecules did not alter the Q-band 
(690 nm). (b) Biomolecular conjugation of VP on DSPE-PEG and 16:0 Lyso PC regained 
fluorescence intensity of VP under physiological relevant condition. Fluorescence 
quenching was observed in Cetuximab-VP as in free VP. (c) Photoactivity is defined as the 
maximal fluoresce intensity (FI) of PSBM in PBS divided by the maximal fluoresce 
intensity (FI) of PSBM in DMSO. (n = 6, ***P < 0.001, n.s.: nonsignificant). (d) SOSG 
reports 1O2 production from photo-activated (690 nm, 10 J/cm2; 10 mW/cm2) PSBMs and 
free VP in PBS at a fixed VP concentration of 5 ?M. (n = 3, *P < 0.05, **P < 0.01, ***P 
< 0.001). N=6, where N equals the number of wells assessed over 3 trials. 
 
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5.3.3 Photodestruction of Glioblastoma Cells Using PSBMs 
In the above studies, we have shown that PSBMs could improve the 
photoactivity of VP. We next investigated if this improvement in photoactivity 
translates to an enhancement in phototoxicity in vitro using two EGFR-positive GBM 
cell lines (U251, U87) (Figure 5.4). Here, 16:0 Lyso PC-VP, DSPE-PEG-VP, 
Cetuximab-VP, and free VP were incubated with GBM cells at different concentrations 
(0-3 ?M) for 24 hours prior to 690 nm light activation (20 J/cm2, 150 mW/cm2, Figure 
5.4a). 16:0 Lyso PC-VP, DSPE-PEG-VP, and free VP all showed negligible dark 
toxicity in U251 or U87 cells (Figure 5.4b-d). It is not surprising that Cetuximab-VP 
(above 0.5 ?M) showed a modest 25% dark toxicity in U251 cells (Figure 5.4b), as it 
has been verified that conjugation of 7 VP molecules onto Cetuximab does not take 
away the targeting functionality of Cetuximab [276, 278]. At 24 hours following PDT, 
the viability of cancer cells was assessed by MTT assay. Free VP was found to be the 
most potent PDT agent with an IC50 of 0.44 ?M?J/cm
2 in U251 cells (Figure 5.4a), 
despite having the least retention of photoactivity and the lowest singlet oxygen yield 
under physiological conditions (Figure 5.3c,d). A significant decrease in phototoxicity 
was observed with 16:0 Lyso PC-VP (IC50: 1.68 ?M?J/cm
2), Cetuximab-VP (IC50: 8 
?M?J/cm2), and DSPE-PEG-VP (IC50: >60 ?M?J/cm
2) (Figure 5.4a,c). This 
remarkable variation in the photo-cytotoxicity of PSBMs was also observed in the U87 
cell line (Figure 5.4d). At a fixed PDT dose of 5 ?M?J/cm2, free VP reduced the 
viability of U87 cells by over 95%, and the loss of cell viability was dramatically 
reduced to approximately 40%, 13%, and 2% when using 16:0 Lyso PC-VP, 
Cetuximab-VP, and DSPE-PEG-VP, respectively (Figure 5.4d). This difference in 
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PDT efficacy does not correlate with the photoactivity and singlet oxygen yields of 
PSBMs in physiologically relevant buffers. We showed that DSPE-PEG-VP is the most 
photoactive formulation in a cell-free system, but it has the poorest in vitro PDT 
Figure 6.4 Near-infrared Light Activation of PSBMs Leads to Phototoxicity of 
Glioblastoma cells. (a) U251 glioblastoma cells were incubated with PSBMs at 
different VP concentrations (0-3 ?M) for 24 hours prior to light illumination (0 or 20 
J/cm2, 150 mW/cm2). Cell viability was evaluated via MTT assay at 24 hours after 
light activation. Light activated PSBMs (+hv) resulted in a significant loss of U251 
viability compared to (b) PSBM treatments without light activation (?hv). (c) U251 
and (d) U87 glioblastoma cells were incubated with i) free VP, ii) 16:0 Lyso PC-VP, 
iii) DSPE-PEG-VP, or iv) Cetuximab-VP at 37 ?C for 24 hours prior to light 
irradiation at a fixed PDT dose of 5.0 ?M ? J/cm2 (i.e., 0.25 ?M VP and 20 J/cm2). 
(n > 3, *P < 0.05, **P < 0.01, ***P < 0.001). N?3, where N equals the number of 
wells assessed over 3 trials. 
95 
 
 
efficacy against GBM cells. These results motivated us to evaluate the cellular uptake 
and location of PSBMs. 
 
5.3.4 PSBM Uptake 
We compared the uptake of free VP and PSBMs in U87 and U251 cells at different 
drug incubation concentrations (0-1 ?M). After 24 hours of incubation in U87 cells, 
free VP showed the highest percentage of cellular uptake at 0.36?0.08% per ?g of 
protein, compared to 16:0 Lyso PC-VP uptake at 0.13?0.02% per ?g of protein. Both 
DSPE-PEG-VP and Cetuximab-VP had the lowest uptake percentage in U87 cells at 
0.06?0.01% per ?g of protein and 0.07?0.0134% per ?g of protein, respectively 
(Figure 5.5a). Similarly, the reduced cellular uptake of PSBMs was observed in U251 
cells (Figure 5.5b). Comparative analyses showed comparable PSBM uptake between 
the U87 and U251 cells at the concentrations used (Figure 5.6). It is likely that the 
stealth function of PEG, or the overall change in amphipathic structure, inhibited the 
cellular uptake of DSPE-PEG-VP [29, 284]. Additionally, we have also reported that 
the limited intracellular uptake of Cetuximab-VP is due to the low photosensitizers-to-
antibody ratio of Cetuximab-VP required for maintaining the antibody?s targeting 
ability, as well as the limited amount of EGFR molecules presented on the cancer cell 
surface [278]. The cellular uptake of VP (or PSBM) was further correlated with PDT-
mediated viability reduction to determine the PDT efficiency (i.e., percent of cell 
viability reduction per fmole of intracellular VP). Interestingly, the PDT efficiency of 
16:0 Lyso PC-VP (~2.2%) was found to be significantly higher than that of free VP 
(~1.1%), DSPE-PEG-VP (0.1-0.3%), and Cetuximab-VP (0.6-0.9%) in both U87 and 
96 
 
 
U251 cell lines (Table 5.1). These results suggest that the cytotoxic mechanisms of 
PSBM-PDT are likely to be different from that of VP-PDT. 
Light activation of photosensitizers not only result in cytotoxicity but can also 
generate fluorescence signal from the relaxation of excited singlet-state 
photosensitizers back to the ground state. Recently, Gliolan? (ALA) has been 
approved for use to visualize malignant tissue during glioma surgery [285]. This use of 
ALA-induced fluorescent protoporphyrin IX (PpIX) to accurately define brain tumor 
margins is critical to achieving optimal surgical outcomes [286]. We next tested if 
PSBMs at 0.25 ?M can be used for fluorescence imaging of U251 cancer cells (Figure 
5.5c). After 24 hours incubation at 37?C and followed by washing to remove loosely 
attached or non-attached agents, live cells treated with free VP showed the highest VP 
fluorescence signal, compared to cells treated with 16:0 Lyso PC-VP, DSPE-PEG-VP, 
or Cetuximab-VP (Figure 5.5c). Quantification of the fluorescence images showed that 
free VP exhibited ~2-10 fold higher fluorescence signal in U251 cells, compared to 
using 16:0 Lyso PC-VP, DSPE-PEG-VP, or Cetuximab-VP (Figure 5.7). While it is 
clear that free VP uptake is greater than that of the PSBMs, extraction data reveals that 
uptake is not significantly different for 16:0 Lyso PC-VP, DSPE-PEG-VP, and 
Cetuximab-VP at the fixed incubation concentration of 0.25 ?M. However, fluorescent 
imaging quantification shows 16:0 Lyso PC-VP is slightly higher than DSPE-PEG-VP 
and Cetuximab-VP. It is well established that fluorescent probes can respond to 
changes in their microenvironments (e.g., pH) [287, 288], which possibly explains this 
observation. While fluorescent imaging is warranted, the more accurate method is 
extraction due to VP being solubilized under the same conditions. It is obvious that our 
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ability to effectively image cancer cells correlates with the amount of the 
photosensitizing agents associated with the cells. However, these data do not suggest 
that 16:0 Lyso PC-VP, Cetuximab-VP, and DSPE-PEG-VP are poor imaging agents, 
as signal-to-background ratio of tumor to the surrounding normal tissue is a more 
important parameter to be carefully investigated in vivo. 
Figure 6.5 Cellular Uptake of PSBMs and Free VP. Quantification of (a) U87 and (b) U251 
intracellular VP concentrations at 24 hours post-incubation with free VP, 16:0 Lyso PC-VP, 
DESP-PEG-VP, and Cetuximab-VP (0-1 ?M) using extraction method. (c) Comparison of 
uptake of PSBM and free VP at 0.25 ?M in both U87 and U251 cell lines. (n = 5, *P < 0.05). 
(d) Representative fluorescence imaging of VP photosensitizer (red) in U251 cells incubated 
with 0.25 ?M free VP or PSBMs for 24 hours. Nuclear staining (blue, Hoechst); Scale bar = 
50 ?m. N?3, where N equals the number of wells assessed over 3 trials. 
 
 
98 
 
 
Figure 6.6 Comparison of PSBMs and Free VP Uptake in GBM Cell Lines. 
Quantification of (a) Free VP, (b) 16:0 Lyso PC-VP, (c) DSPE-PEG-VP, and (d) 
Cetuximab-VP intracellular VP concentrations at 24 hours post-incubation, in U251 and 
U87 cells, using extraction method. (n = 5, n.s.: nonsignificant). 
 
 
 
 
Table 6.1 PDT Efficiency of Free VP and PSBMs in Glioblastoma Cells. 
Groups* U251 U87
(PDT efficiency** ) (PDT efficiency** )
Free VP 1.12?0.00% 1.07?0.00%
16:0 Lyso PC-VP 2.30?0.03% 2.14?0.29%
DSPE-PEG-BPD 0.29?0.09% 0.09?0.08%
Cet-VP 0.94?0.28% 0.57?0.24%
*Drug incubation concentration: 0.25 ?M VP equivalent. **PDT efficiency:
percent of viability reduction per fmole of intracellular VP. PDT drug-light
interval: 24 hours. PDT light dose: 690 nm, 20 J/cm2, 150 mW/cm2.
 
99 
 
 
Figure 6.7 Quantification of VP Fluorescence Signal. U251 Cells were incubated with 
Free VP, 16:0 Lyso PC-VP, DESP-PEG-VP, and Cetuximab-VP (0.25 ?M) for 24 hours. 
Quantification of fluorescence signal as determined by ImageJ software. (n = 4, *P < 
0.05, ***P < 0.001, n.s.: nonsignificant). 
 
5.3.5 Altered Subcellular Localization of PSBM  
One of the preferred sites of localization for VP photosensitizer is mitochondria 
[138, 139]. Activation of VP by light causes photodynamic disruption of the 
mitochondrial membrane, which triggers the release of cytochrome c, a potent initiator 
of apoptotic cell death. PDT has also been shown to photochemically degrade the anti-
apoptotic protein Bcl-2, while leaving pro-apoptotic proteins Bad and Bax unaffected 
[138, 139]. This shifts the balance in target cells from an anti-apoptotic state to a more 
pro-apoptotic state. To determine whether PSBMs are localized to the mitochondria, 
we incubated U251 cells with PSBMs for 24 hours and treated cells with MitoTracker 
prior to examination using a confocal fluorescence microscope (Figure 5.8a). U251 
cells that had been incubated with free VP and Cetuximab-VP exhibited a strong and 
diffuse fluorescence signal throughout the cells. Converging the VP image and 
MitoTracker image gave a pseudo-colored yellow fluorescent pattern (Figure 5.8a), 
and the profile plot analyses confirmed the localization of free VP and Cetuximab-VP 
100 
 
 
at mitochondria (Figure 5.8b). In contrast, 16:0 Lyso PC-VP clustered in the peri-
nuclear region with a reduced degree of mitochondria localization in U251 cells 
(Figure 5.8). The DSPE-PEG-VP fluorescence signal in U251 cells was minimal and 
heterogeneously distributed across different cells (Figure 5.8). It was further confirmed 
that peri-nuclear staining of 16:0 Lyso PC-VP related more to lysosomal incorporation 
(Figure 5.9) compared to mitochondria similar to that observed by others [203]. 
101 
 
 
Figure 6.8 Mitochondrial Localization of Free VP and PSBMs in U251 Cells. (a) 
Representative confocal fluorescence images of U251 cells obtained after 24 hours of 
incubation with free VP, 16:0 Lyso PC-VP, DSPE-PEG-VP, and Cetuximab-VP. VP signal 
pseudo-colored in red. Mitochondria were stained with 100 nM MitoTracker (M7514; pseudo-
colored in green); merging of images where yellow indicates co-localization of VP (red) and 
mitochondria (green). Nuclear staining (Hoechst; pseudo-colored in blue). Scale bar = 20 ?m. 
(b) Profile plots were created using ImageJ software showing co-localization of mitochondria 
and free VP (or PSBM). N ? 3. 
 
 
 
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Figure 6.9 Lysosomal Localization of Free VP and PSBMs in U251 Cells. (a) 
Representative confocal fluorescence images of U251 cells obtained after 24 hours 
of incubation with free VP, 16:0 Lyso PC-VP, DSPE-PEG-VP, and Cetuximab-VP. 
VP signal pseudo-colored in red. Lysosomes were stained with 100 nM of 
LysoTracker probes (DND-99; pseudo-colored in green); merging of images where 
yellow indicates co-localization of VP (red) and lysosomes (green). Nuclear staining 
(Hoechst; pseudo-colored in blue). Scale bar = 20 ?m. (b) Profile plots were created 
using ImageJ software showing co-localization of lysosome and free VP (or PSBM). 
N ? 3. 
We next investigate the ability of PSBM-based PDT to trigger apoptotic cell death, 
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damage lysosomes, regulate autophagosome formation, and depolarize mitochondrial 
membrane potential (??m). To account for the variations in PSBM uptake, U251 cells 
were incubated with different concentrations of free VP (0.15 ?M), 16:0 Lyso PC-VP 
(0.25 ?M), DSPE-PEG-VP (0.50 ?M), or Cetuximab-VP (0.50 ?M) to achieve an 
equivalent intracellular VP concentration (~0.35 ?moles per gram of protein) prior to 
light irradiation. In order to compare the degree of which the PSBMs can induce 
apoptosis, flow cytometric detection of apoptosis with Annexin V and propidium 
iodide was performed. Light fluence was fixed at 20 J/cm2, which has been shown to 
reduce cell viability to nearly 0% at 24 hours after treatment (Figure 5.4; MTT assay). 
For flow cytometry-based apoptosis detection, cells were examined at 4 hours after 
light exposure to evaluate acute cellular injury (Figure 5.10a). Free VP, 16:0 Lyso PC-
VP, and Cetuximab-VP all induced similar levels of apoptotic (~20%) and dead cells 
(~11%) (Figure 5.10a), while DSPE-PEG-VP had little impact on cell viability (~96%) 
although having the same intracellular concentration. Due to lysosomal localization, 
PDT using 16:0 Lyso PC-VP induced a high degree of lysosomal membrane 
permeabilization in U251 cells, decreasing the mean fluorescent intensity of 
LysoTracker to 40% compared to that of the no treatment control (Figure 5.10b). PDT 
treatment with free VP, DSPE-PEG-VP, and Cetuximab-VP modestly reduced the 
mean fluorescent intensity of LysoTracker (Figure 5.10b), suggesting minimal 
lysosomal damage. A number of studies have already shown that PDT-mediated 
lysosomal damage could induce autophagy [289-292]. Here, we evaluated 
autophagosome formation at 6 and 24 hours after PDT using free VP or PSBMs. 
Immunoblotting results indicated that PDT using free VP and PSBMs all induced an 
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increase of LC3-II in comparison to the no treatment control at 24 hours post-treatment 
(Figure 5.10c). Increase in LC3-II indicates the accumulation of autophagosomes, 
which can be seen even at 6 hours after irradiation for free VP, 16:0 Lyso PC-VP, and 
Cetuximab-VP. However, this does not guarantee autophagic degradation [293]. 
Owing to mitochondrial localization (Figure 5.8), PDT using free VP or Cetuximab-
VP induced a high level of ??m depolarization in U251 cells (Figure 5.10d). No ??m 
depolarization was observed using 16:0 Lyso PC-VP and DSPE-PEG-VP for PDT of 
U251 cells (Figure 5.10d). A potent mitochondrial oxidative phosphorylation 
uncoupler, FCCP, was used as a positive control for ??m depolarization [294]. 
Considering the time of incubation used (i.e., 24 hours) in our study and above findings, 
16:0 Lyso PC-VP co-localization with MitoTracker (Figure 5.9b) could potentially be 
associated with auto-lysosomes containing mitochondria as suggested in Martins? 
study [295]. The observed Cetuximab-VP and free VP co-localization with 
LysoTracker could also be associated with auto-lysosomes containing mitochondria 
(Figure 5.9b). Further investigations into the cell/patient line-dependent phototoxicity 
and autophagy-associated cell death of PSBMs are warranted. 
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Figure 6.10 Acute Cellular Injury, Lysosomal Membrane Permeabilization, 
Autophagosome Formation, Mitochondrial Membrane Depolarization, and 
Combination Treatment using PSBMs and Free VP. Cells were incubated with 
PSBMs at their respective uptake concentrations (i.e., 0.15 ?M of free VP, 0.25 ?M 
of 16:0 Lyso PC-VP, 0.5 ?M of DSPE-PEG-VP, and 0.5 ?M of Cetuximab-VP) for 
24 hours prior to PDT. (a) Flow cytometric detection of apoptosis with Annexin V 
FITC and propidium iodide was performed at 4 hours after light irradiation (20 J/cm2, 
150 mW/cm2) to detect living (Q4), early apoptotic (Q3), late apoptotic (Q2), and 
dead cells (Q1). (b) Lysosomal membrane permeabilization was assessed 30 minutes 
after PDT. (c) Immunoblotting of LC3-II was assessed 6 and 24 hours after PDT. (d) 
Mitochondrial membrane depolarization was assessed immediately after PDT. (e-f) 
Cell viability was determined via MTT assay at 24 hours post-PDT (690 nm, 0.1 
J/cm2 for U251; 0.1 and 1 J/cm2 for GBM39, 10 mW/cm2, bottom illumination). (n > 
3, *P < 0.05, **P < 0.01, ***P < 0.001. 
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Studies by Martins et al., Coincotta et al., and Kessel showed that PDT directed 
against both lysosomes and mitochondria could enhance the overall efficacy of photo-
killing of malignant human cells, murine hepatoma cells and sarcoma [295-299]. Some 
of the earlier studies utilized different types of photosensitizers for lysosomal targeting 
(e.g., NPe6, EtNBs) and mitochondrial targeting (e.g., VP), and thus require different 
wavelengths of light for PDT activation (i.e., NPe6, ?max at 630nm; EtNBs, ?max at 652 
nm; VP ?max at 690 nm). While PDT with photosensitizers accumulating in 
mitochondria leads to loss of ??m and the release of pro-apoptotic mediators (e.g., 
cytochrome c), light activation of photosensitizers that localizes in lysosomes could 
induce cell death via the release of proteolytic lysosomal enzymes [300]. Rizvi et al. 
have shown that 690 nm light action of Visudyne? and lipid-anchored Liposomal VP 
enhances PDT outcomes via multi-organelle targeting [299]. Similarly, here, we 
showed that single wavelength (i.e., 690 nm) light activation of both free VP and 16:0 
Lyso PC-VP, which target mitochondria and lysosomes, respectively, could achieve a 
significant reduction in viability at 72.6?5.1%, compared to the modest viability 
reduction achieved by VP-PDT (43.8?1.8%) or 16:0 Lyso PC-VP-PDT (30.9?6.9%) 
(Figure 5.10e). Similarly, PDT using a mixture of VP and 16:0 Lyso PC-VP 
significantly reduced the viability of GBM39 cells by ~80%, compared to that using 
free VP (15.7?11.9% viability reduction) or 16:0 Lyso PC-VP (58.9?6.9% viability 
reduction) alone (Figure 5.10f). In contrast to that observed in U87 and U251 cell lines, 
16:0 Lyso PC-VP was found to be more phototoxic compared to free VP in GBM39 
cells at a fluence of 0.1 J/cm2 (Figure 5.10f). Thus, further mechanistic investigations 
into the cell/patient line-dependent phototoxicity of PSBMs are needed. We speculate 
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that GBM39 cells showed resistance to free VP effect could be due to the activation of 
pro-survival mitophagy to maintain cell homeostasis. Thus, by compromising 
lysosomal function, as done using 16:0 Lyso PC-VP, GBM39 cells become more 
sensitive to photodamage triggered by VP-PDT. Martins and others have already 
showed this concept [295]. At a higher fluence of 1.0 J/cm2, both 16:0 Lyso PC-VP 
and free VP led to GBM39 killing of approximately 80%, while the combination killed 
virtually 100% of the GBM39 cells. While the combinations of free VP and 16:0 Lyso 
PC-VP led to a greater decrease in cell viability compared to monotherapy, a 
synergistic effect (Two-way ANOVA, interaction P value < 0.0001, GraphPad Prism) 
was only seen within the GBM39 cells at 1.0 J/cm2. A follow up full dose?response 
matrix study is required to identify and quantify synergistic, additive, and antagonistic 
dose regions. Additional animal research is also warranted to examine the benefits of 
combining 16:0 Lyso PC-VP and free VP to enhance the overall PDT efficacy. 
5.4 Conclusion 
In this study, we synthesized a panel of VP-based PSBMs with different molecular 
weights and physiochemical properties. We systematically analyzed how different 
classes of biomolecules (i.e., lipid-, PEG-, and antibody-based) influence the physical, 
photochemical, and biological properties of the VP photosensitizers. The simple and 
reproducible formulation strategies did not alter the Q-band of VP for PDT using 690 
nm light activation. The instability of free VP in physiologically relevant media led to 
a complete fluorescence quenching. Conjugation of VP to 16:0 Lyso PC or DSPE-PEG 
slightly improved VP?s fluorescence emission by up to 20%. Static quenching of VP 
molecules on cetuximab was expected, and could be de-quenched upon lysosomal 
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proteolysis of Cetuximab-VP for PDT, confirming previously findings [276, 278]. For 
all PSBMs and free VP, a strong correlation between the degree of photoactivity and 
singlet oxygen yield was observed in a cell-free system. However, no correlation was 
found between the evaluated physicochemical properties and the photodynamic 
therapeutic index of PSBMs in two GBM cell lines (U87, U251). 
The uptake kinetics and subcellular localization of PSBMs were found to be the 
more critical factors for effective PDT and fluorescence imaging. Despite lower singlet 
oxygen production, the 16:0 Lyso PC-VP exhibited enhanced delivery of VP into GBM 
cells, compared to DSPE-PEG and Cetuximab-VP, and could be visualized by 
fluorescence imaging. While the clathrin?mediated endocytosis pathway and lysosomal 
proteolysis mechanism of Cetuximab-VP in cancer cells have been well-established by 
us and others, here, we present the new evidence that proteolyzed Cetuximab-VP co-
localizes to mitochondria to efficiently induce mitochondrial membrane (??m) 
depolarization upon light activation in GBM cells. Light activation of 16:0 Lyso PC-
VP or DSPE-PEG-VP did not induce ??m depolarization. This can be explained by 
the localization of 16:0 Lyso PC-VP in lysosomes, as well as the heterogeneous and 
negligible uptake of DSPE-PEG-VP in GBM cells. Our findings suggest that 
engineering PSBM for effective PDT is enabled by close examination of cellular 
trafficking and subcellular localization in order to predict their effectiveness. 
Importantly, it is apparent that no single parameter is predictive. Compared to PSBMs, 
free VP was found to be a more potent PDT agent in vitro due to its higher cellular 
uptake and mitochondrial incorporation, which leads to a pronounced mitochondrial 
oxidative stress with apoptosis activation. 
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It is well accepted that subcellular localization of the photosensitizer plays a 
significant role in the type of cell death mechanism that predominates. Employing 
different types of photosensitizers or functionalizing photosensitizers with targeting 
moieties to target one or multiple subcellular organelles (e.g., mitochondria, lysosome, 
endoplasmic reticulum, golgi, plasma membrane) have also been reported [296, 297]. 
Here, we verified that 16:0 Lyso PC redirects VP to lysosomes. More importantly, we 
demonstrated that single wavelength light activation of both free VP and 16:0 Lyso PC-
VP targeting multiple subcellular organelles is a superior approach compared to using 
free VP or 16:0 Lyso PC-VP alone. Above insights of the molecular and cytotoxic 
mechanisms of PSBMs are critical to inform the design, prediction and evaluation of 
more sophisticated photo-nanomedicine for glioma PDT and imaging. 
In summary, our results reveal the variable trafficking and cytotoxicity of clinically 
relevant PSBMs for PDT, providing important insights into methods of PSBM 
evaluation as well as strategies to select PSBMs based on cellular uptake, subcellular 
localization, and cytotoxic mechanisms. While biologically informed combinations of 
PSBMs and free-form photosensitizers may lead to enhanced therapeutic effects, the 
cellular uptake of PSBMs need to be enhanced to fully exploit the dual targeting of 
organelles. Chapter 6 will discuss the positive and the negative impacts of lipids (i.e., 
liposomes, lipid nanoparticle) on drug delivery, and highlight the numerous parameters 
that need to be considered for liposome design.  
 
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7. Chapter 6. Immunological and Toxicological 
Considerations for Liposome Use4 
6.1 Introduction 
Liposomes are vesicular structures composed of one or more concentric lipid 
bilayers surrounding an aqueous cavity [301-303]. The bilayers are predominantly 
composed of phospholipids, where the polar head groups interface with the outer and 
inner aqueous phases and the hydrophilic tails are sequestered within the bilayer [303]. 
Since their discovery by Alec Bangham in the 1960s [304, 305], liposomes have been 
studied extensively as drug delivery vehicles due to their capacity to load both 
hydrophilic and hydrophobic agents, as well as their high biocompatibility and tunable 
size, charge, and surface properties [301-303]. Liposomal encapsulated drugs first 
reached the clinic in 1995 with the FDA-approval of Doxil? (Liposomal doxorubicin) 
for the treatment of AIDS-related Kaposi's sarcoma, and Doxil? was later approved to 
treat ovarian cancer, and multiple myeloma [306]. Since the FDA approval of Doxil?, 
numerous liposomal formulations have been employed in the clinic for a wide array of 
applications, including cancer therapeutics, fungal disease treatment, analgesics, PDT, 
and viral vaccine delivery [190, 246]. However, despite the increasing prominence of 
liposomal drugs in the clinic, there is still limited knowledge regarding their 
toxicological effects on healthy cells and tissues, as well as the immunological 
responses they can elicit. 
                                                 
4 This chapter was adapted (modified) from Inglut C.T., et al., ?Immunological and Toxicological 
Considerations for the Design of Liposomes?. Nanomaterials 2020, 10(2), 190. No special 
permission is required to reuse this article published by MDPI under the terms and conditions of 
the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/). 
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Phospholipids, the primary building blocks of liposomes, are amphipathic 
molecules, meaning they have a hydrophilic region (e.g., polar phosphate head) and a 
hydrophobic section (e.g., non-polar fatty acid tail). When hydrated in an aqueous 
solution under artificial conditions, phospholipids spontaneously organize into 
liposomes due to their thermodynamic phase properties and self-assembling 
characteristics [307]. The physio-biochemical characteristics of liposomes can be 
modified by altering the types and ratios of phospholipids, as well as incorporating 
cholesterol into the bilayer and decorating the liposomal surface with PEG. These 
modifications can have drastic effects on healthy cells and tissues, as well as activate 
or suppress the immune system. These complex interactions, therefore, have immense 
implications for the clinical use of liposomal formulations and will be discussed in 
depth later in this review. Extensive research has been done to develop a variety of 
techniques to achieve optimized liposome formation and drug loading. Incorporating 
therapeutic agents into liposomes can be achieved either during liposome formation 
(e.g., passive loading) or after liposome formation (e.g., active loading). Passive 
loading can be further divided into three categories: mechanical dispersion methods, 
solvent dispersion methods, and detergent removal methods [301, 308, 309]. 
Alternatively, active loading can be accomplished by establishing a pH gradient, 
causing the unionized drugs that penetrate the lipid bilayer to become ionized due to 
the low pH within the liposome, resulting in entrapment [310, 311]. FDA approval has 
been granted for both passively loaded liposomal agents (e.g., Visudyne? and 
AmBisome?) and actively loaded liposomal agents (e.g., Doxil?, Myocet?, and 
Onivyde?) [312]. Most of the clinically used liposome-based products are 
112 
 
 
administered by intravenous (IV) injection, though some are also given by 
intramuscular injection (e.g., Inflexal? V and Epaxal?), by epidural injection (e.g., 
DepoDur?), or by intrathecal injection (e.g., Depocyt?) [246]. 
Liposomes are particularly useful for delivering hydrophobic agents, which 
otherwise have poor solubility in aqueous solutions and limited bioavailability [24, 25]. 
VP (also known as benzoporphyrin derivative), for example, is a hydrophobic 
photosensitizer that is used for PDT, a light-based therapeutic modality. While VP self-
aggregates in aqueous solutions, Liposomal VP (marketed as Visudyne?) has 
improved solubility for intravenous administration and is FDA-approved to treat wet 
age-related macular degeneration [145, 191]. To date, Visudyne? is being evaluated in 
multiple clinical trials for PDT of cancer due in part to its favorable pharmacokinetic 
profiles (i.e., rapid clearance), leading to low phototoxic skin reactions [313]. In 
addition to improving drug solubility, liposomes have a variety of other advantages as 
drug delivery vehicles, such as high biocompatibility, modifiable physicochemical and 
biophysical properties, and protection of large drug payloads from degradation, 
inactivation, and dilution in circulation [302, 314]. Liposomes are also being 
engineered to deliver drugs to parts of the body that are exceptionally difficult to access 
(e.g., central nervous system) [315]. Intravenously-injected liposomal drug 
formulations provide the additional benefit of preferentially accumulating in tumor 
tissues due to the enhanced permeability and retention effect, thereby incorporating an 
inherent mechanism for passive tumor targeting [27, 28].  
Another key advantage is that liposomal encapsulation of drugs increases the 
therapeutic index by decreasing the accumulation of drugs in organs and other healthy 
113 
 
 
tissues [26-28]. For example, anthracyclines, a group of potent chemotherapeutics, can 
induce severe cardiotoxicities, including cardiomyopathy and congestive heart failure. 
This limits the range of doses that can be safely administered. Using liposomes to 
deliver doxorubicin, one of the most widely used and most cardiotoxic anthracyclines, 
avoids drug accumulation in myocardial tissues, improving the overall response rates 
and attenuating the risk of cardiotoxicity [316, 317]. For instance, liposomal 
encapsulation of doxorubicin increased the median lethal dose (LD50) by approximately 
2-fold in mice (32 vs. 17 mg/kg) and 1.5-fold in beagle dogs (2.25 vs. 1.5 mg/kg) 
compared to free-form doxorubicin [318]. Similarly, liposomal encapsulation of 
irinotecan (i.e., Onivyde?), another chemotherapeutic agent used in multiple solid 
organ cancers, enables improved pharmacokinetics, increased maximum tolerable 
dose, and protection of irinotecan from hydrolysis-mediated inactivation in circulation 
[319]. Though liposomes have demonstrated the ability to reduce the accumulation of 
drugs in some organs and healthy tissues, they tend to reroute them to other sites. One 
meta-analysis concluded that on average, less than 1 % of nanoparticles end up 
delivered to solid tumors [320]. This underscores a key problem for liposomal agents, 
which largely accumulate in organs associated with the mononuclear phagocyte system 
(MPS), where they can impair liver function by attenuating phagocytic capacity and 
depleting macrophages [321]. 
While liposomes are used to reduce systemic toxicity from the encapsulated agents, 
the liposome itself can impart toxicity to normal tissues and elicit an immune response 
(Figure 6.1). Cationic liposomes, which are largely studied for gene delivery, are 
known to elicit toxicity in macrophages, macrophage-like cells, and monocyte-like 
114 
 
 
cells, as well as decrease the secretion of important immuno-modulators [322-324]. 
Liposomes can also trigger immunogenic responses, though the nature and degree of 
these responses can vary depending on several properties, including surface charge, 
size, and pegylation [325-327]. Therefore, despite the many advantages of using 
liposomes for drug delivery, there are also important drawbacks that must be 
considered, as they present real challenges for clinical translation. This mini-review 
will explore these limitations in-depth, with an emphasis on toxicities and unfavorable 
interactions with the immune system, to shed light on some of the lesser-known areas 
where liposomes have room for improvement as drug delivery vehicles. 
Figure 7.1 Schematic of a Liposome and its Common Building Blocks. The Liposomal 
bilayer is composed of phospholipids (cationic or anionic) and cholesterol, and the surface can 
be decorated with polyethylene glycol (PEG). Hydrophilic drugs can be entrapped within the 
aqueous core, while hydrophobic drugs can be loaded into the lipid bilayer. Once injected into 
the bloodstream, a protein corona, comprised largely of apolipoproteins, immunoglobulins, and 
complement proteins, is formed on the liposome surface. The protein corona, which is impacted 
by the Liposomal surface chemistry, governs liposome-cell interactions. The immunological 
and toxicological effects caused by each liposome component, imparted on cells throughout 
the body, are summarized throughout the diagram. 
  
115 
 
 
6.2 Toxicological Evaluation of Liposomes and Their Building Blocks 
Liposomes are typically composed of numerous building blocks, including 
phospholipids, cholesterol, and PEG, each impacting their functionality. 
Phospholipids, the main component of liposomes, are diverse molecules with a variety 
of isomers. The most commonly used phospholipids for liposome formation include 
lecithins from egg and soy, as well as sphingomyelins [328]. Common variations to the 
phospholipid hydrophobic acyl (fatty acid) tail include the degree of saturation (i.e. 
saturated vs unsaturated) and length, which can impact the oxidizing properties and 
drug release profiles of liposomes [329]. For example, Luo et al. showed that PDT-
mediated oxidation of unsaturated lipids (i.e., 1,2-dioleoyl-sn-glycero-3-
phosphocholine, or DOPC, 18:1 PC) in the liposomal bilayer could facilitate the 
controlled release of doxorubicin. Increasing the concentration of DOPC in liposomes 
from 0% to 10% enhanced the release rate of doxorubicin from liposomes by up to 10-
fold upon light-activation of photosensitizers [330]. Different phospholipid head 
groups, with some of the most prevalent being phosphatidylcholine (PC), 
phosphatidylserine (PS) and phosphatidylethanolamine (PE), can alter the surface 
charge and the polymorphic phase (i.e., the type of assembly) in an aqueous medium 
[331]. Although PC is a neutral molecule, it has a positively charged choline group 
attached to a negatively charged phosphate group, making it a zwitterionic molecule. 
Alternatively, PS contains a negatively charged phosphate group attached to the serine 
group, giving it an overall negative charge, mainly due to the low (2.2-3.6) pKa of the 
carboxyl group within the serine [332]. More recently, there is an increased research 
interest in formulations of lipid-drug (or photosensitizer) conjugates to improve the 
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pharmacokinetics and therapeutic index of liposomal therapeutics [196, 333, 334]. 
Similarly, the incorporation of PEG, a polymeric steric stabilizer, is widely used to 
increase the blood circulation time of liposomal formulations [29, 284]. The addition 
of PEG into Liposomal doxorubicin extended the terminal half-life from 16.4 hours to 
77 hours in patients with metastatic breast cancer [335, 336]. Once liposomes enter the 
bloodstream, plasma proteins interact with and adsorb onto the liposome surface. It has 
been previously shown that the liposomal surface chemistry (e.g., charge and 
pegylation) impacts the blood protein adsorption (known as the ?protein corona"), 
which influences how liposomes interact with healthy and diseased cells [337-339]. 
Protein adsorption occurs due to an advantageous increase in entropy, resulting in a 
decrease in Gibbs free energy (?Gabs) [340]. A comprehensive study by 
Hadjidemetriou et al. examined the in vitro and in vivo protein corona formation on 
three different types of liposomes, including pegylated liposomes, non-pegylated 
liposomes, and monoclonal antibody-conjugated liposomes [341]. In this study, 
pegylated liposomes had the lowest protein adsorption at ~100,00 ?g protein/?M lipid, 
while non-pegylated liposomes had double the amount of protein adsorption. However, 
all liposomes had relatively consistent protein corona compositions, with the most 
abundant bound proteins being apolipoproteins, immunoglobulins, and complement 
proteins.   
While the topic of nanoparticle-protein adsorption is an active field of study [340-
343], several studies have evaluated the toxicity and side effects of liposomes. 
Papahadjopoulos et al. first reported the cytotoxicity of liposomes in healthy mouse 
fibroblast L929 cells in 1973. They discovered that incorporation of 3% or 10% of 
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lysophosphatidylcholine (lysolecithin, lysoPC) into liposomes increases cellular 
uptake (i.e., polykaryocytosis) by ~7%. The number of viable L929 cells after 
liposomal incubation was significantly lowered from 97% to less than 37% [344]. 
Similarly, Pagano et al. showed that extensive accumulation of egg yolk lecithin 
vesicles in Chinese hamster lung fibroblast V79 cells results in considerable cell death 
[345]. One of the first studies evaluating liposomal toxicity in vivo was carried out by 
Adams and colleagues in 1977 [346]. In this study, five different types of liposomes 
with varying net surface charges were synthesized and injected into the mouse brain. 
Results showed that liposomes containing 9 mol% of dicetyl phosphate (net negative 
charge) were the most toxic, causing epileptic seizures and rapid death in animals. 
Injection of dicetyl phosphate alone resulted in the death of all mice within one hour. 
Brain sections stained with haematoxylin and eosin (H&E) revealed tissue necrosis, 
nerve damage, and loss of Nissl substance. Similarly, liposomes containing 9 mol% of 
stearylamine (net positive charge) caused widespread brain damage and respiratory 
failure. Alternatively, liposomes containing 9 mol% of phosphatidic acid (net negative 
charge) were well tolerated with only small hemorrhages and necrosis limited to the 
injection site. The least toxic liposomes were those composed of 45 mol% of lecithin 
or dipalmitoyl lecithin (neutral charge), which resulted in minimal toxic reactions and 
morphological changes, even after 3 repeated doses. While liposomes are typically 
considered pharmacologically inactive with minimal toxicity [301, 314], their toxicity 
is tightly related to the type of model, exposure time, dose, and/or surface properties. 
For example, Shi et al. showed the LD50 using F98 glioma cells was 6.07 ?M for empty 
DPPC-based cationic liposomes while DSPC-based anionic liposomes had an LC50 > 
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509 ?M. Within an intracranial F98 glioma rat model, carboplatin loaded cationic 
pegylated liposomes resulted in a median survival time of 29 days, while anionic 
pegylated liposomes had a median survival of 49.5 days [347]. Similarly, Fan et al. 
showed that cationic liposomes incubated with bone marrow derived dendritic cells had 
an LC50 around 0.2 mg/ml, while ovalbumin-loaded anionic liposomes had an LC50 > 
4 mg/ml [348]. In Section 6.2, we examine some of the different mechanisms through 
which liposomes induce cellular toxicity.  
As liposomes rose in popularity, many research groups began to modify the 
phospholipid structure and test different surface chemistries [349, 350]. Studies in the 
early 1990s by Parnham et al. [351] and Lappalainen et al. [352] examined and 
suggested different assays for the screening of liposomal toxicity. Some of these in 
vitro viability and morphology assays include MTT, 3H thymidine uptake, and lactate 
dehydrogenase. It was shown that 40 ?M of liposomes containing dimethyldioctadecyl 
ammonium bromide (18:0) and dioleoylphosphatidylethanolamine (DOPE, 18:1 PE) 
reduce the viability of human cervical squamous cell carcinoma cells by 50% [352]. 
Liposomes containing N-(2,3-Dioleoyloxy-1-propyl)trimethylammonium methyl 
sulfate (DOTAP, 18:1 TAP) at a concentration of 10 ?M or 40 ?M only had minor 
effects on the cancer cells [352]. There is also evidence demonstrating anionic lipids 
have a higher thrombogenic potential [353]. Anionic liposomes have been shown to 
induce platelet aggregation in guinea pigs [354], as well as blood cell aggregation in 
mice, which became temporally trapped in lung capillaries [355]. To ensure that 
intravenous injection of liposomes does not induce pro-aggregatory effects or 
pharmacodynamic changes (e.g. necrosis, edema, loss of tissue) at the site of entry, 
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appropriate platelet aggregation assays and local tolerance studies should be 
performed. The mechanical damage during injection should be accounted for as well, 
and distinguished from any toxicological effects [351]. In April 2018, the FDA 
published updated guideline for liposome products in industry [356]. Although explicit 
details are not specified, critical parameters ranging from physicochemical 
characterization (e.g., pegylation, net charge, leakage rate during shelf storage) to 
pharmacokinetic profiles are established. Radiolabeled mass-balance excretion and 
metabolism studies are required, as they reveal quantitative connections to toxicity, and 
provide information on the in vivo activity of new chemical entities including 
pharmacology and pharmacokinetics [357]. 
6.2.1 Toxicity of Cationic Liposomes 
Cationic liposomes have been studied extensively to enhance the stability and 
delivery of negatively charged nucleic acid therapeutics [358-363], and their toxicities 
and side effects have also been carefully examined [322-324, 364-371]. For example, 
Filion and Phillips studied the toxicity and immunomodulation of seven different 
DOPE (18:1 PE)-based cationic liposomes that range from +15.0 to +42.9 mV [322]. 
In this comprehensive study, liposomes containing DOTAP, dimethyldioctadecyl 
ammonium bromide, 1,2-dimyristoyl-3-trimethylammonium-propane (DMTAP, 14:0 
TAP), 1,2-dipalmitoyl-3-trimethylammonium-propane (DPTAP, 16:0 TAP), or 1,2-
stearoyl-3-trimethylammonium-propane (DSTAP, 18:0 TAP) were highly toxic to 
macrophages and monocyte-like U937 cells, but not to resting or activated T 
lymphocytes. It was speculated that these cationic liposomes were non-toxic to T cells 
due, in part, to minimal uptake. Additionally, the cytotoxicity of cationic liposomes 
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increased at higher zeta potentials. Dimethyldioctadecyl ammonium bromide -
containing liposomes (+40.0 ? 9.0 mV) were the most toxic to macrophages, with a 
50% effective dose (ED50) <10 nmol/ml, while DSTAP-containing liposomes (+15.0 ? 
6.3 mV) were the least toxic, with an ED50 >1000 nmol/ml. Loading plasmid DNA into 
liposomes significantly reduced the zeta potential, but did not decrease their 
cytotoxicity [322]. The cytotoxicity of the liposomes was significantly lowered by 
incorporating 10 mol% of 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (DPPE, 
16:0 PE)-PEG2000 or replacing DOPE with 1,2-dipalmitoyl-sn-glycero-3-
phosphocholine (DPPC, 16:0 PC). These modifications reduced the binding and 
endocytosis of liposomes by ~96% by macrophages and hindered their ability to 
destabilize the endosomal membrane [322]. In addition to inducing cytotoxicity in 
macrophages, the cationic liposomes also mitigated the secretion of nitric oxide and 
tumor necrosis factor-?, which are important signaling molecules for triggering non-
specific defense against pathogens and enhancing T and B lymphocyte responsiveness 
[322].  
Similarly, Takano et al. and Aramaki et al. showed that the intracellular uptake of 
cationic liposomes (~42 mV) in mouse macrophage-like cell line RAW264.7 correlates 
with the degree of apoptosis and the generation of ROS [323, 324]. However, the 
incorporation of 7.5% and 10% DPPE-PEG into liposomes reduced their uptake by 
~30-45-fold and increased cell viability by ~20-38%. Inclusion of DPPE-PEG also led 
to a decrease in ROS production [323], which diminished the toxic effects of liposomes 
[324]. Soenen et al. showed that adverse effects from cationic liposomes can be reduced 
by using ROS scavengers or a calcium channel blocker in 3T3 fibroblasts [372]. Based 
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on these studies and many others  [322, 365-367, 369-371, 373], cytotoxicity could be 
readily detected in a wide range of cells upon increasing the zeta potential values of 
liposomes beyond ~30 mV, in a charge dependent manner. In general, cationic 
liposomes are uptaken by phagocytic cells to a greater extent and induce the 
formulation of ROS, which damages organelles (e.g., mitochondria), and promotes 
higher intracellular Na+ levels, compared to neutral and negatively charged liposomes 
[323, 324, 371-375].  
Using animal models, Kedmi et al. showed that intravenous injection of cationic 
liposomes (+54.3 ? 6.1 mV) containing DOTAP-induced hepatotoxicity and pro-
inflammatory responses [365]. The cationic liposomes significantly reduced the body 
weight of C57BL/6 mice by up to 5.5% and increased liver enzymes in serum by 3-6-
fold compared to neutral and negatively charged (?59.2 ? 4.9 mV) liposomes. Cationic 
liposomes also increased the amount of Th1 and Th17 cytokines by 10-24 fold at 2 
hours post-injection [365]. In rats, Knudsen and colleagues demonstrated that the 
administration of DOTAP/CHOL liposomes (+53 ? 2 mV) led to an upregulation of 
oxidative stress response gene and the DNA repair enzyme in the liver, as well as 
increased DNA strand breaks in the lungs [366]. While cationic liposomes are 
promising vehicles for gene delivery, understanding their toxicities to immune cells 
and regulation of inflammatory responses are critical for improving therapeutic benefits 
and avoiding immune toxicity.  
6.2.2 Interactions with the Mononuclear Phagocyte System 
While liposomes improve the pharmacokinetic profiles of therapeutics, they also 
redirect the therapeutic payloads to the MPS (previously known as the 
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reticuloendothelial system, RES), [376-378]. The MPS is a collection of phagocytic 
cells (e.g., progenitor, monocytes and macrophages) located within the bone marrow, 
lymph nodes, spleen, and liver. The MPS is responsible for the removal of antigens 
from the body, and is the main site of liposome accumulation [379, 380]. Several rodent 
studies have shown that Doxil? accumulates within the liver and spleen (e.g., the two 
major organs of the MPS [376]) to a greater extent than other organs shortly after 
intravenous injections [381-385]. In cynomolgus monkeys, Kimelberg et al. found that 
liposomal encapsulation of [3H]MTX caused a 160-fold increase in [3H]MTX uptake 
by the spleen compared to the free-form [3H]MTX [378]. It has been observed by many 
researchers that the redirection of liposomal chemotherapy to the MPS induces toxicity 
and depletion of immune cells, especially the Kupffer cells in the liver [321, 386-391]. 
To study the impact of liposomes on the phagocytic function of MPS cells in mice, 
Allen et al. tested several compositions of egg PC-CHOL liposomes [386]. Injection 
(IV) of larger, multilamellar liposomes (synthesized using 200-450 nm filters) 
significantly decreased the phagocytic index of the liver by approximately 50% in mice, 
compared to using small unilamellar liposomes. The phagocytic index did not recover 
until 3 weeks post-injection, and further doses of liposomal injections lead to 
accumulated in the spleen. Multiple injections caused a significant decrease in the 
phagocytic index of the spleen and a 2.5-fold increase in the spleen weight. The 
function and weight of the spleen did not recover until 8 days after the last liposome 
injection. 
Several groups have also studied the impact of drug-containing liposomes on liver 
macrophages. Daemen et al. showed that intravenous injection of non-pegylated, short-
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circulating Liposomal doxorubicin in rats reduced the number and phagocytic capacity 
of liver macrophages by up to 85-90%, compared to placebo liposomes [321]. The 
phagocytic capacity of liver macrophages was impaired for 8 days, and full recovery 
took 2 weeks. A follow-up study examined the effects of long-circulating Liposomal 
doxorubicin containing ~4.8 mol% PE-PEG on the phagocytic activity of macrophages 
[391]. The pegylated liposomes reduced the phagocytic capacity and number of liver 
macrophages by 63% at 72 hours after injection. Compared to non-pegylated 
liposomes, pegylated liposomes delayed the cytotoxic effects on liver macrophages by 
24 hours and modestly reduced the depletion of macrophage by 7%. While pegylation 
of liposomes reduces the blood protein adsorption and increases blood circulation time, 
the ultimate impact of pegylated and non-pegylated liposomes on macrophages and 
liver function was similar. 
Research groups have also shown that within the tumor microenvironment, 
liposomes can polarize tumor-associated macrophages (TAMs) [392-394]. The 
reprogramming of classically activated (M1) TAMs, which have a role in eradicating 
tumor cells in their early stages, to alternatively activated M2-like TAMs have been 
shown to promote tumor growth and metastasis, as shown in Figure 6.2 [395, 396]. In 
an orthotopic ovarian carcinoma (ID8- vascular endothelial growth factor -GFP) mouse 
model, La-Beck et al. showed that empty liposomes increased tumor volume by over 
3-fold and doubled the number of metastatic sites, compared to the vehicle control 
[392]. Similarly, Sabnani et al. and Rajan et al. found that compared to the vehicle 
control (e.g., 5% dextrose solution, saline), empty pegylated liposomes accelerated the 
growth of subcutaneous TC-1 tumors in mice [394, 397]. The accelerated tumor growth 
124 
 
 
was attributed to diminished macrophage function and number of tumor-specific T cells 
[397], as well as a 66% decrease in M1-TAMs and a 100% increase in M2-TAMs [394]. 
Based on these results, strategies to overcome immunosuppression and tumor 
progression triggered by liposomes may potentially improve the efficacy of some 
liposomal anticancer drugs. New insights into the underlying molecular mechanisms 
may yield the key to designing new liposome-based immunotherapies.  
Figure 7.2 Intravenously Injected Liposomes Impact Cancer Progression. Liposomal 
polarization of classically activated M1 tumor associated macrophages (TAMs) to M2-like 
TAMs can result in carrier-induced immunosuppression and accelerated tumor development. 
Compared to vehicle control, empty (placebo) liposomes (a) accelerated tumor growth in a TC-
1 subcutaneous mouse model and (b) increased the number of metastatic sites in an 
orthotopically implanted ID8-VEGF-GFP ovarian carcinoma mouse model on day 36. 
Reprinted from Inglut C.T., et al. and La-Beck et al., Harnessing Liposome Interactions With 
the Immune System for the Next Breakthrough in Cancer Drug Delivery. Front. Pharmacol., 
12 March 2019. (No permission required, under the Creative Commons Attribution License, to 
use, distribution or reproduction in other forums.) 
 
6.2.3 Effect of Cholesterol Content on the Toxicity of Liposomes 
Cholesterol is commonly used to alter the mechanical properties and functionality 
of liposomes [398-400]. Cholesterol improves the stability and drug entrapment 
efficacy of liposomes by enabling more dense assembly of phospholipids [398-403]. 
Smaby et al. demonstrated that cholesterol condenses the lipids (e.g. from 63 to 44 
?2/molecule; 16:0-18:1 PC liposomes), thereby increasing the compressibility of 
125 
 
 
liposomes from 123 to 401 mN/m [404]. The concentration of cholesterol can also 
impact the surface charge and permeability of liposomes [401]. Incorporation of 50 
mol% cholesterol into 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC, 18:0 PC) 
liposomes reduced the zeta potential from 2 to -5.5 mV [405]. The presence of 
cholesterol also reduced the permeability coefficients of phospholipid vesicles to small 
molecules and ions [406]. A previous study showed that cholesterol prevents the 
aggregation and fusion of liposomes, leading to enhanced penetration into the skin 
[403]. In additional to structural stability and drug efflux related studies, several groups 
have examined the toxicity and immunogenicity of cholesterol. Adams et al. showed 
that injection of 5-10 mg of cholesterol into the mouse brain resulted in mild signs of 
edema and loss of internal neuronal structures [346]. Others have shown that free-form 
cholesterol induces macrophage apoptosis via the stiffening of the endoplasmic 
reticulum [407] and promotes the production of tumor necrosis factor-? and interleukin 
6 (IL-6) [408]. Roerdink et al. showed that incorporation of 50 mol% of cholesterol 
into DSPC-based liposomes decreased their liver accumulation (from 70% to less than 
40%) and extended the blood circulation half-life by 3-fold in rats [409]. It has also 
been shown that cholesterol-containing liposomes decrease macrophage uptake and 
immunoglobulin G response in mice [410]. Similarly, incorporation of high levels (43 
mol%) of cholesterol in ovalbumin-liposomes significantly reduced immunoglobulin 
G response by 10-fold in mice [411]. In a Li 210 leukemia mouse model, Ganapathi et 
al. showed that cytarabine-loaded liposomes containing 50 mol% of cholesterol 
increased animal survival by up to ~20%, compared to using cytarabine-loaded 
liposomes containing 10 mol% of cholesterol or free-form cytarabine [412]. However, 
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it is unclear if this enhanced cancer killing is due to increased cellular uptake of 
liposomes or an increase in the sustained release of cytarabine from cholesterol-
containing liposomes. While the incorporation of cholesterol into liposomes mitigates 
macrophage uptake and antibody production, other studies have shown that cholesterol 
could activate the immune system by promoting interactions with C3, a protein 
associated with the complement system [413-415]. This is discussed extensively in 
Section 6.3.2. 
 
6.3 Activation of the Immune System 
Biomaterials introduced into the body can initiate immune responses in the host 
tissue. As shown in previous work, host defense mechanisms and inflammatory 
responses to biomaterials are known to be activated by the complement response [416]. 
The products from complement response activation may assist neutrophil aggregation 
and migration towards inflammatory sites [416]. Earlier research suggests that PEG-
based hydrogels cause increased inflammatory responses compared to a silicone-based 
hydrogels. This change was visualized through increased IL-1? expression in the host 
tissues after implantation of a PEG-based hydrogel [417]. It is also thought that 
complement activation can induce a foreign body reaction [416]. During this process, 
host inflammatory cells and foreign body giant cells are activated to destroy the 
implanted material. Previous studies have demonstrated that thick macrophage 
capsules accumulate on PEG-based hydrogels as a result of the FBR when compared 
to other hydrogels lacking PEG [417]. 
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Liposomes, like other nanoparticles used in drug delivery, interact with proteins in 
the body to trigger innate immune system responses [418-420]. Once liposomes have 
been administered to the patient, circulating proteins can adsorb to the surface of the 
liposome, thereby creating a protein corona unique to the characteristics of the 
liposome. The protein corona can then induce the activation or suppression of various 
immune responses [421]. Particularly, liposomes interact with complement proteins to 
activate the complement cascade and increase the body?s response to antigens [418, 
421, 422]. The incorporation of PEG into liposomes has been shown to promote 
immunogenic responses via complement activation and the anti-PEG antibody 
production [37]. For these reasons, liposomes are being used as vaccine adjuvants to 
potentiate the immunological response to antigens. Antigen-liposomal complexes can 
increase and maintain exposure of the antigen in the lymph nodes, allowing for 
enhanced uptake by immature phagocytic antigen-presenting cells (e.g. macrophages 
and dendritic cells) [423-425]. A notable antigen-liposome adjuvant system, AS01, has 
been studied in multiple different disease clinical trials [426], and was FDA approved 
as a shingles vaccine (Shingrix) in 2017.  
The liposomal surface charge and the size can impact the type and efficiency of 
the immune response. A study by Badiee et al. investigated the effects of the surface 
charge of recombinant rgp63-loaded liposomes on the immune response of mice with 
leishmania after subcutaneous injections. Neutral liposomes were found to promote a 
Th1 (e.g., interferon ?, IL-12) immune response more efficiently than positively 
charged liposomes. The largest IL-4, immunoglobulin Ga, and immunoglobulin G1 
response were induced by positively charged liposomes. A Th2 (e.g. IL-5, IL-6, IL-10) 
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response, but not a Th1 immune response, was caused by liposomes with a negative 
surface charge [427]. In regards to liposome size, a follow-up study by Badiee et al. 
showed that 100 nm liposomes induce a Th2 response, while liposomes larger than 400 
nm induce a Th1 response [428]. This was due, in part, to their altered pharmaceutics 
and intracellular trafficking. Similarly, Henriksen-Lacey et al. showed that 
intramuscularly injected ~200 nm CAF01 cationic liposomes induced the greatest IL-
10 response in mice [429], while medium (500 nm) liposomes induced the largest 
interferon-? and IL-1? response. The 200 nm and 1.5 ?m liposomes were drained more 
rapidly by the lymph node, while the 500 nm liposomes were well retained at the site 
of injection [429]. 
6.3.1 Pro-Inflammatory Cytokine Production 
Liposomal interactions with blood cells lead to cytokine production and the 
activation of the immune system [360, 365, 430, 431]. The activation of the 
complement cascade also promotes the secretion of cytokines from immune cells [432, 
433]. This concept is well demonstrated in the first liposomal gene silencing clinical 
trial (NCT00882180). Liposomes were loaded with siRNAs that were chemically 
modified to minimize immunostimulation and administered intravenously. However, 
there was still a dose-dependent induction of proinflammatory cytokines. Cytokines, 
including IL-6, IP-10, granulocyte colony-stimulating factor, and tumor necrosis 
factor-?, peaked 6 hours post-injection. One of the largest liposome doses, 1.25 mg/kg, 
resulted in the most IL-6 present in the patients? blood (~9000 pg/mL). There was also 
a ~3-10 fold increase in Bb complement protein for patients who received a dose larger 
than 0.2 mg/kg [360]. Yamamoto et al. demonstrated that cytokine induction by HEPC-
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based liposomes is comparable to that of a potent activator of monocyte and 
macrophages, lipopolysaccharides. Also, incubation of liposomes with human 
peripheral blood induced monocyte-derived cytokines (IL-6, IL-10, IL-1?, interferon-
y and tumor necrosis factor-?) but not lymphocyte-derived cytokines (IL-2, IL-4 and 
IL-5). In the same study, it was shown that the size of liposomes directly affects the 
degree of cytokine release, in vitro, in a size-dependent manner. Liposomes with an 
800 nm diameter induced the production of 6000 pg/mL of IL-1?, while 50 nm 
liposomes produced virtually none [430]. 
For gene delivery using liposome-DNA complexes, previous reports have shown 
that nonspecific cytokine production from toll-like receptor-9 positive immune cells 
occurs due to unmethylated CpG dinucleotides (CpG motifs) abundantly present in 
pDNA [434, 435]. Liposome-DNA complexes also induced the production of type I 
interferons, irrespective of the frequency of CpG motifs in DNA and the expression of 
toll-like receptor-9  [436]. However, a study conducted by Yasuda et al. showed that 
immune activation by liposome-DNA complexes is highly dependent on the type of 
cationic liposome. In murine macrophage-like cell line, RAW264.7, non-CpG 
containing Lipofectamine2000 liposomes (+35.1?mV) induced the largest cytokine 
production (500-900 pg/ml), while DOTMA/DOPE and DOTMA/CHOL liposomes 
(+32.7 mV) produced less than one-fifth of that amount [437]. Furthermore, 
Lipofectamine2000 liposomes containing non-CpG motif DNA induced interferon-? 
and IL-6 production by macrophages cultured from toll-like receptor-9 deficient mice 
[437]. These liposome-induced inflammatory responses are critical to consider for 
130 
 
 
clinical applications, as increased levels of cytokines can cause side effects (e.g., grade 
1?2 chills/rigors, flu-like symptoms) [360, 438]. 
6.3.2 Recognition of Liposomes via Complement Activation 
The complement system is a part of the innate immune system responsible for the 
recognition of foreign objects in the body [432]. It is composed of over 30 plasma proteins that 
interact with each other to opsonize (e.g., C3b) and clear liposomes, as well as induce a series 
of inflammatory responses by liberating anaphylatoxins (e.g., C3a, C4a, C5a) (Figure 6.3) 
[422, 432]. Early reports indicated that liposomes activate the alternative pathway via C3 
binding and conversion, which is independent of antibody recognition. Cunningham et al. 
showed that C3 conversion occurs when cationic liposomes are incubated in serum from C1r-
deficient and C2-deficient patients [414]. Liposomes without cationic lipids or with less than 
20 mol% of cholesterol were unable to convert C3 in C2-deficient serum [414]. However, later 
reports show complement activation is dependent on the surface chemistry and composition of 
liposomes [439-441]. A study by Chonn and colleagues showed that liposomes containing 20 
mol% of anionic lipids activated the classical pathway in a Ca(2+) dependent manner. Calcium 
is required in the classical pathway, as it is used for C1qr2s2 complex formation. Cationic 
liposomes activated the alternative pathway by reducing the C3-C9 levels in Ca(2+) depleted 
serum. In addition, liposomes containing unsaturated lipids or 45 mol% of cholesterol 
promoted complement protein interactions. In contrast, neutral liposomes, even at a high lipid 
concentration of 50 mM, did not activate the complement pathways [415]. Bradley et al. 
showed anionic liposomes can also activate the classical pathway in an antibody-independent 
manner via direct binding of C1q to the liposomal surface [441]. Liposome size, pegylation, 
and cholesterol content can also contribute to the degree and nature of complement activation. 
For example, studies have suggested that larger liposomes, over 200 nm in diameter, were 
associated with an increase in complement protein opsonization and alternative pathway 
131 
 
 
activation [442, 443]. Liposomes containing PEG typically induce IgM antibody production, 
followed by (classical) complement activation [37]. A study by Alving et al. showed a 
cholesterol dependent activation, with liposomes containing 73 mol% cholesterol promoting 
the greatest classical pathway activation [444]. While examining the hypersensitivity reactions 
(HSRs) induced by different components of Doxil?, Szebeni et al. demonstrated that 
negatively charged PEG-PE is a potent C protein activator, and the degree of complement 
activation correlates with HSRs [445]. When PEG-PE-based liposomes were incubated with 
human serum, there was a 2.5-4-fold increase in soluble complement S5b-9 (C5b-9), compared 
to HSPC-based liposomes which had no impact on complement activation [445]. Activation of 
C5b-9 following exposure to pegylated liposomes has been associated with development of 
HSR [446], which is discussed in the following section.  
132 
 
 
Figure 7.3 Schematic Overview of the Complement System Highlighting Two of the Main 
Activation Pathways (Classical vs. Alternative). Previous studies have shown that classical 
pathway activation is initiated by antibody-liposome binding, pegylated liposomes, and anionic 
liposomes. The alternative pathway can be initiated by cationic liposomes, liposomes 
containing more than 40 mol% cholesterol (Chol), and liposomes larger than 200 nm in 
diameter. During the complement cascade, C3b opsonin covalently binds to the surface of the 
liposome, marking it for removal by the MPS. The released anaphylatoxins (C3a, C4a, and 
C5a) prompt the activation of leukocytes. 
 
6.3.3 Hypersensitivity Reactions 
           Liposomes have also been shown to induce HSR [447-454]. HSR are non-IgE-
mediated pseudo-allergic immune responses, which usually develop immediately after 
intravenous infusion when the body is exposed to an antigen or liposomes [446]. While 
6.8% of the 705 patients who received Doxil for refractory AIDS-related Kaposi?s 
sarcoma experienced HSR with symptoms ranging from dyspnea and tachypnea to 
133 
 
 
hypotension and hypertension, the underlying causes remain unclear [454, 455], with 
some researchers attributing similar reactions to complement activation. [456, 457]. 
Later reports have shown HSR are correlated with PEG-induced complement 
activation, however high levels of C activation may be necessary, suggesting the 
involvement of other pathogenic factors [445, 446, 458]. This subset of HSR, known 
as C activation-related pseudoallergy, can be reduced by slowing the rate of infusion, 
diluting the Doxil dose, or premedicating with a corticosteroid [459]. In a 4-patient 
study, grade 3 HSR induced by Doxil occurred almost immediately after the start of 
infusion in all patients and treatment was stopped. Premedication with ranitidine and 
hydroxyzine prior to the resumption of Doxil infusion eliminated HSR in 3 of the 4 
patients [450]. Similarly, data analysis performed by Chanan-Khan et al. showed, on 
average, 8% of people who received Doxil experienced HSR [446]. Additionally, 3% 
of refractory ovarian cancer patients who were pretreated with corticosteroids and 
antihistamines to minimize adverse reactions still experienced HSR. Chanan-Khan et 
al. further showed HSR occurrence can be as high as 45% for patients receiving Doxil. 
Within this Phase 1 clinical study, 92% of the patients who experienced HSR had 
significantly elevated Plasma SC5b-9 levels [446]. HSR have been suggested to be 
caused by the liposomal vehicle of Doxil rather than the encapsulated drug, as these 
reactions are not known to occur with standard doxorubicin [445, 446]. It should also 
be noted that other chemotherapies, such as Taxol (paclitaxel), which relies on a 
formulation vehicle, and carboplatin, are known to cause HSR [460, 461]. 
Using pig models, Szebeni et al. showed that pulmonary hypertension reactions 
are dependent on composition, size, and administration method of liposomes [449]. 
134 
 
 
Large, neutral, multilamellar liposomes composed of 1,2-dimyristoyl-sn-glycero-3-
phosphocholine (DMPC, 14:0 PC) and DSPC induced pulmonary hypertension, while 
smaller liposomes (< 200 nm) caused no hemodynamic changes [449]. An additional 
study by Szebeni et al. showed that highly negatively changed liposomes containing L-
?-phosphatidylglycerol initiated CARPA to the greatest extent in pigs, suggesting that 
surface charge may also play a role [462]. F?l?p et al. showed a 300-600% increase in 
pulmonary arterial blood pressure after the infusion of pegylated Liposomal 
prednisolone sodium phosphate in pigs [448]. The risk of these side effects was reduced 
by using a slow infusion protocol (0.04?ml/kg/h) with a 3-step dose escalation [448]. It 
is now recommended that Doxil be administered at 1 mg/min to help minimize these 
adverse reactions [463]. Within a 2018 Nature Nanotechnology Perspective, authors 
suggested that the next steps for minimizing HSR and CARPA are identifying induced 
biomarkers and understanding the underlying mechanisms via in vitro assays [464].  
 
6.3.4 Anti-PEG Response 
Liposomal interactions with immune cells can cause antibody production against 
liposome components, such as PEG [37, 326, 465-469]. The rapid production of 
antibodies (e.g. immunoglobulin G, M, E) from B lymphocytes, in a T cell-independent 
manner, is another mechanism in the recognition and removal of liposomes from the 
body [470]. Antibody binding impacts the blood circulation time and treatment efficacy 
of liposomes and initiates classical complement activation. IgM antibodies are the first 
produced, by the spleen, in response to an antigen or liposomes, as seen in Figure 6.4 
[467, 471, 472]. One of the first studies to notice an increase in antibody production 
135 
 
 
was by Bucke and colleagues in 1998 when studying the biodistribution of liposomes 
with different surface modifications [473]. Additionally, Cheng et al. were one of the 
first to show that anti-PEG IgM monoclonal antibodies were generated by the spleen 
of mice in response to PEG-immunoconjugates [474]. Based on blood concentration 
levels and binding, IgM antibodies were more efficient in clearing the PEG-
immunoconjugates from the blood than immunoglobulin G [474]. 
Since then, several groups have studied antibody production in preclinical animal 
models, which helped to explain the accelerated blood clearance seen after repeated 
dosing of liposomes. Ishida et al. showed that pegylated liposomes incubated with 
serum derived from rats pre-injected with liposomes had an increased liposome-protein 
binding index by approximately 2-fold compared to serum obtained from rats that were 
not pre-injected with liposomes [326]. Also, there was a 17-fold increase in IgM 
binding to pegylated liposomes compared to the non-pegylated liposomes. A follow-
up study by Ishida and colleagues revealed a T cell-independent response to pegylated-
liposomes in both Wistar rats and BALB/c nu/nu mice [465]. A relatively low anti-
PEG immunoglobulin G response was seen 3-5 days after injection of pegylated 
liposomes and a potent IgM response occurred 3-10 days post injections in Wistar rats. 
In the absence of T-cells, a ~9-fold increase in IgM was observed 10 days post-injection 
in BALB/c nu/nu mice [465]. Additionally, using a pig model, Kozma et al. showed 
that anti-PEG IgM concentration in blood peaks at 7-9 days after low dose intravenous 
injection of pegylated liposomes [458]. The anti-PEG IgM levels remained elevated 
above normal, by approximately 10-fold, for 6 weeks. Using ex vivo spleen cells 
derived from pigs pre-injected with pegylated liposomes, they showed an increase in 
136 
 
 
IgM+ B-cells bound to Doxil? from 1.88% to 5.50% [458]. The association between 
the underlying mechanism of PEG immunogenicity and the ?accelerated blood 
clearance? of subsequent liposome injections will be discussed in the next section. In 
contrast to the immunogenic effects, it is critical to point out that PEG may have 
additional immunosuppressive effects by decreasing inflammation and fibrosis, as seen 
in organ preservation [475], which warrants further investigation.  
Figure 7.4 Schematic Overview of the Events that Alter the Pharmacokinetics of 
Repeated Intravenous Doses of Pegylated Liposomes. The first dose of pegylated liposomes 
stimulates the production of anti-PEG IgM antibodies by B cells within the spleen. This leads 
to the accelerated blood clearance of subsequent doses, during which circulating anti-PEG 
IgMs bind to the liposomes, initiating classical complement activation and decreasing blood 
circulation half-life.  
6.3.5 Accelerated Blood Clearance 
As previously mentioned, the addition of PEG to liposomes can increase their 
blood circulation half-life and prolong the uptake by the MPS [29]. It is also well 
accepted that after repeated injections of pegylated liposomes, they lose their ?stealth? 
ability and are more quickly cleared from the blood [30-43]. This phenomenon, known 
as ?accelerated blood clearance? (ABC), is caused by the abundant secretion of anti-
PEG immunoglobulin M, G, and E, followed by the opsonization of C protein 
fragments (complement activation), and finally uptake by macrophages, depicted in 
Figure 6.4 [30-42]. Dams et al. first observed a dramatic change in the circulatory half-
137 
 
 
life of intravenously injected 99mTc-labeled pegylated-liposomes in rats and monkeys, 
but not mice [30]. In a male rhesus monkey, the blood circulation half-life of the second 
dose of pegylated-liposomes dropped from 87.5 hours to 14.2 hours. Although further 
liposome injections after the second dose had similar kinetics as the first, the ABC of 
the second dose occurred if it was administered 1-4 weeks after the first. Rats transfused 
with blood or serum from rats that were previously injected with liposomes also 
experienced ABC. The percent of the injected dose of liposomes in blood transfused 
rats at 4 hours dropped from 52% to less than 23% in the blood, suggesting the ABC is 
caused by a soluble serum factor [30].  
Studying the ABC of liposomal chemotherapy in rodents may not be ideal, as 
Laverman et al. confirmed that Doxil? injections in rats lead to hepatosplenic 
macrophage depletion, and therefore a second injection of Doxil? did not experience 
ABC [31]. However, if the first dose was empty pegylated liposomes, an injection of 
Doxil? within one week was cleared rapidly from circulation in rats [31]. Similarly, in 
Beagle dogs, Suzuki et al. showed that repeated doses of Doxil? at 20 mg/m2 did not 
have altered pharmaceutics, but repeated doses less than 2 mg/m2 were more rapidly 
cleared from the blood and elicited an IgM response [40]. The half-life of the second 
injection at 2 mg/m2 was reduced from 24.1 hours to 1.5 hours. Larger doses of Doxil? 
could have caused damage to B cells within the spleen, attenuating antibody 
production, or the uptake capacity of Kupffer cells may have been saturated/suppressed 
by the first large dose [40]. It is critical to point that the clinical relevance of ABC 
phenomenon induced by repeated injection of pegylated liposomes remains debatable, 
since Gabizon et al. observed a significant decrease in clearance (P < 0.0001) from the 
138 
 
 
1st through the 3rd cycle of pegylated Liposomal doxorubicin in humans [476]. The 
wide use of PEG in healthcare, hygiene, and beauty products suggests that most patients 
will likely have pre-existing anti-PEG antibodies [477], which could potentially impact 
the degree of ABC in patients. Other parameters that have been shown to impact the 
ABC of liposomes are lipid dose, with increasing amounts of the prior dose altering the 
pharmacokinetics of the subsequent injections in a sigmoid manner [34], liposome 
composition, with unsaturated lipids causing a more pronounced ABC [33], and time- 
and frequency- of injections [30, 32, 33, 478]. 
6.4 Conclusion 
For two and a half decades, liposomal drug formulations have been administered 
in the clinic for the treatment of a variety of ailments, ranging from cancer to fungal 
disease. They boast an array of advantageous features, including biocompatibility, 
tunable properties, and capacity for loading hydrophilic and hydrophobic agents, 
making them convenient drug delivery vehicles. However, despite their clinical 
relevance and therapeutic potential, there is still a scarcity of knowledge regarding the 
downsides associated with the administration of liposomes. This mini-review presents 
a summary of existing knowledge regarding such limitations, divided into two main 
sections: 1.) Toxicological Evaluation of liposomes and their Building Blocks, and 2) 
Activation of the Immune System. One of the main toxicological concerns is that 
cationic liposomes, which are primarily used for nucleic acid delivery, can be toxic to 
macrophages and reduce their secretion of important immunomodulators. Additionally, 
following intravenous injection, liposomes end up sequestered in the organs of the 
MPS, such as the spleen and liver. This induces toxicity in these organs, causing the 
139 
 
 
depletion of cells that are critical for proper immune system function. As for activation 
of the immune system, liposomes can induce an inflammatory response that is 
characterized by the release of pro-inflammatory, monocyte-derived cytokines. 
Similarly, liposomes have been shown to activate the complement pathway, though the 
degree of activation is dependent on several factors such as size, charge, and mol% of 
cholesterol. Another important mechanism of immune activation is the anti-PEG 
response, which occurs when pegylated liposomal formulations activate the production 
of PEG antibodies by B lymphocytes. This leads to recognition and clearance by the 
immune system. In fact, repeated administrations of pegylated liposomes have been 
shown to cause accelerated blood clearance, robbing liposomes of their stealth 
capabilities. This underscores the idea that the dosing of liposomal agents must be 
carefully optimized to avoid premature clearance and maximize the therapeutic benefits 
of each injection. Depending on injection speed, liposomes can also induce 
hypersensitivity reactions, including cardiopulmonary distress. Mode of delivery and 
speed of administration are therefore key considerations for clinical applications of 
liposomal formulations. Overall, liposome-mediated toxicity to healthy tissues, 
coupled with the activation of the immune system, raises important concerns regarding 
their use. While liposomes are an attractive tool to reduce the dose-limiting side effects 
of therapeutics in patients, the liposomes themselves can impart new toxicities that are 
still not fully understood. There is still much work to be done to understand the 
mechanisms through which liposomes interact with the immune system before the full 
potential of liposome-based systems for gene and drug delivery can be achieved. 
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8. Chapter 7. Carrier-free, Amorphous Verteporfin 
Nanoparticles for Enhanced Photodynamic Cancer 
Therapy and Brain Drug Delivery 
7.1 Introduction 
As mentioned in Section 2.3.2, verteporfin (VP, also known as benzoporphyrin 
derivative) is a photosensitizer with excellent singlet oxygen yield for PDT [152] and 
minimal systemic toxicities due to rapid clearance primarily via liver-mediated 
mechanisms [313]. PDT involves the intravenous administration of VP followed by 
non-thermal 690 nm laser activation of VP to generate highly cytotoxic, short-lived 
ROS. While VP has proven advantageous in a wide range of disease contexts, the 
molecule is relatively hydrophobic [479] and, to date, has required carrier systems for 
clinical delivery [264, 480, 481]. In 2000, FDA approved a liposomal formulation of 
VP (Visudyne?) for PDT of age-related macular degeneration [482, 483]. Significant 
clinical efforts have since demonstrated the utility of Liposomal VP for PDT of 
chorioretinal conditions [484] and cancers [485, 486]. 
Liposomes have improved the solubility of photosensitizers [487], but cell-level 
delivery can still be augmented. For example, studies have shown that free-form VP 
accumulated within cancer cells by 1-2 orders of magnitude higher than Liposomal VP 
formulations [196, 488, 489]. Furthermore, lipophilic photosensitizers are generally 
encapsulated in the lipid bilayer of liposome with limited loading capacity and 
increased risk of photosensitizer self-quenching?a bottleneck for an effective PDT in 
the clinic. Lovell and colleagues demonstrated that up to 80,000 porphyrin-lipid 
conjugates could be packed into the lipid bilayer of each liposome [203]. The high 
packing density results in highly self-quenched porphyrins that convert absorbed light 
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into heat energy rather than releasing fluorescence signal and singlet oxygen 
generation. The reported maximum loading capacity of porphyrin in liposomes is 15 
molar % with up to 11-fold self-quenching. Considering the clinical value of liposomes, 
but understanding their limitations for photosensitizer delivery, we sought to develop 
a carrier-free nanoformulation with efficient photosensitizer delivery and release for 
enhanced PDT. 
Carrier-free nanodrugs offer the opportunity to deliver higher levels of 
hydrophobic photosensitizers. They allow for a theoretical drug loading capacity of 
100% and eliminate the reliance on lipids, polymers, and/or co-solvents for drug 
delivery. Several synthesis schemes, such as media milling and solvent-antisolvent 
precipitation methods, are available for producing carrier-free nanodrugs with 
crystalline or amorphous structures. Nanodrugs with a crystalline character (referred to 
as nanocrystals) have high saturation solubility, and high stability post injection due to 
the minimized Gibbs free energy [490]. Nanocrystals have been leveraged for 
extended-release of antiretroviral therapies [491-493], anticancer drugs [494, 495], and 
other medications [496]. However, in the context of PDT, the potential for prolonged 
photosensitizer bioavailability from nanocrystal could potentially be unfavorable in 
terms of skin phototoxicity. In contrast, amorphous nanodrugs generally possess a 
higher saturation solubility and, consequently, an increased dissolution velocity 
(creation of high Cmax, reduction of tmax) compared to equally sized nanocrystals [497]. 
However, prerequisites for utilization of carrier-free, amorphous photosensitizer 
products for PDT are that the stability can be maintained throughout the shelf-life of 
the product and photosensitizers can be light-activated for ROS generation. 
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Here, we report the first carrier-free, amorphous photosensitizer nanodrug using 
VP (termed NanoVP). The size of the NanoVP can be precisely controlled either by 
adjusting the solvent-to-antisolvent ratio or changing the initial drug concentration. 
NanoVP exhibits excellent stability over 1-year storage, enhances VP accumulation in 
cells, and fully de-quenches in biological systems for superior photochemical singlet 
oxygen generation compared to the standard Liposomal VP formulation. In comparison 
with other photosensitizer formulations that are commonly used in the clinic, we 
demonstrate that NanoVP is a superior PDT agent for the treatment of brain tumors, as 
well as the safe and selective opening of the BBB without damaging normal brain 
tissues. 
7.2 Methods 
7.2.1 NanoVP Synthesis and Characterization 
NanoVP was prepared using the solvent antisolvent precipitation method. Free-
form VP powders (US Pharmacopeia) were dissolved in dimethyl sulfoxide (DMSO, 
solvent) to achieved different initial concentrations (1-40 mM) and then injected 
dropwise into deionized water (antisolvent), under stirring, at room temperature. 
Different solvent-antisolvent ratios (1:50-1:10) were tested for formulating NanoVP. 
Free-form VP molecules not loaded into the NanoVP were removed by dialysis 
(Spectrum Labs, MWCO 300 kDa) against PBS at 4?C for 24 hours. The hydrodynamic 
diameter, polydispersity, and zeta potential were measured using particle sizer and zeta 
potential analyzer (NanoBrook Omni, Brookhaven Instruments). VP concentration was 
determined based on UV-Vis absorbance in DMSO (Synergy Neo2, BioTek 
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Instruments) using established molar extinction coefficient (VP: ? = 80,500 M-1cm-1 at 
435 nm, ?=34,895 M-1cm-1 at 687 nm). Entrapment efficacy is the percentage of VP 
that is successfully entrapped into the NanoVP. Loading capacity is the amount of VP 
loaded per unit weight of the NanoVP. NanoSight tracking analysis (NanoSight LM10, 
Malvern Instruments) was used to determine the number of particles per mL. 
Liposomes are considered ?gold standard? nanocarreirs for VP delivery. Liposomes 
containing VP within phospholipid bilayers were synthesized via freeze-thaw extrusion 
technique [19] and used as a control group. 
7.2.2 Transmission Electron Microscopy 
The size, morphology, and microstructure of NanoVP were studied using a 
transmission electron microscope (TEM; JEOL JEM-2100 LaB6, 200 kV). For 
conventional TEM, NanoVP (10 ?L) was pipetted onto a Lacey carbon grid (Ted Pella) 
and air dried overnight before examination. To exclude the possibility of artifacts due 
to sample dehydration, ionic liquid (3 ?L, Hilem IL-1000, Hitachi) was used as a 
pretreatment reagent for TEM examination of wet specimens. Images were taken at 
high magnifications (10,000-100,000x). Due to small particle sizes and to enhance 
diffraction intensity, selected area electron diffraction was used to collect electron 
diffraction patterns. 
7.2.3 NanoVP Stability, Photoactivity, and Photosensitizer Release Profile 
The stability of NanoVP particles in PBS at 4?C was determined by monitoring 
their hydrodynamic size and polydispersity index using a particle sizer analyzer for up 
to one year. NanoVP singlet oxygen (1O2) generation, self-quenching, and drug release 
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were studied in 96-well plates as described previously [21]. NanoVP and SOSG at 5 
?M were mixed and irradiated with 690 nm light (10 J/cm2, 10 mW/cm2; ML6600, 
Modulight). A multi-mode microplate reader (Synergy Neo2, BioTek) acquired 
fluorescence signals of VP (Excitation/Emission: 435/650-750 nm) or SOSG 
(Excitation/Emission: 504/525 nm) before and after light irradiation. Self-quenching 
(FDSMO/F0) is defined as the fluorescence after disruption of the NanoVP using DMSO 
(FDSMO) divided by the fluorescence of the NanoVP in PBS (F0). Photosensitizer release 
from NanoVP (de-quenching) was studied by monitoring the gain in VP fluorescence 
signal in PBS/DMSO buffer (0-100% v/v) at room temperature or in Eagle?s Minimum 
Essential Medium (Cellgro) with 0-10% v/v fetal bovine serum (FBS, Gibco) at 37?C. 
7.2.4 NanoVP Electrostatic Stabilization 
The electrostatic stability of NanoVP was probed using the neutral polymer, 
poloxamer 407 (Pluronic ? F-127; Sigma), and the cationic polymer, polyethylenimine. 
Polymers were dissolved in water according to manufacturer?s instructions and were 
added to the antisolvent (ultra-pure deionized water) prior to drop wise synthesis or 
after NanoVP particles were formed, at a 1:5 polymer: VP ratio (w:w). In both 
instances, the addition of positively charged polyethylenimine resulted in 
uncontrollable agglomeration, while NanoVP particles coated with poloxamer 407 
appeared to remain stable. Dynamic light scattering and laser Doppler electrophoresis 
measurements showed that NanoVP coated with poloxamer 407 has a hydrodynamic 
diameter of 112.7?5.9 nm (PdI: 0.10?0.01) and a zeta potential of -32.6?1.8 mV. 
ynamic light scattering and laser Doppler electrophoresis measurements cannot be 
accurately performed on large agglomerations.  
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7.2.5 Cancer Cell Cultures 
Human GBM U87 and mouse 3T3 fibroblast cells were obtained from ATCC 
and cultured per vendor?s instructions. The human breast cancer MCF-7 parental cell 
line, the P-gp-overexpressing MCF-7 TX400 subline, and the ABCG2-overexpressing 
MCF-7 MX100 subline were cultured in Eagle's Minimum Essential Medium 
supplemented with 10% FBS, 100 U/mL penicillin, 100 ?g/mL streptomycin, and 0.01 
mg/ml insulin (Sigma) as previously described [196]. Cells were maintained in 5% CO2 
at 37?C and tested to be free of mycoplasma (MycoAlert, Lonza). 
7.2.6 Evaluation of Photosensitizer Uptake and PDT Responses In Vitro 
Cells were cultured overnight in 35-mm Petri dish or 96-well black wall plates (1-
3.3?104 cells/cm2) and then incubated with photosensitizers (i.e., NanoVP, free-form VP, or 
Liposomal VP; 0.25 ?M) for 24 hours. Subsequently, cells were washed twice with PBS and 
incubated with photosensitizer-free complete medium. Photosensitizer uptake in cells was 
determined using extraction methods followed by VP fluorescence measurements 
(Excitation/Emission: 435/700?10 nm, Synergy Neo2, BioTek) or visualized using 
fluorescence imaging (Lionheart, BioTek) as described previously [21]. PDT was performed 
by exposing the cells to 690 nm light (0-10 J/cm2, 10-50 W/cm2, bottom illumination; ML6600, 
Modulight). The generation of intracellular ROS was studied using 2?,7??dichlorofluorescin 
diacetate probe (DCFDA, Thermo Fisher) and the mitochondrial membrane potential was 
examined via TMRE assay (Abcam). Expressions of total and cleaved caspase 3 were examined 
by immunoblotting. At 24 hours after PDT, cell viability was determined using MTT [3-(4,5-
dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] assay or neutral red assay (abcam) 
following the vendor?s protocol. For dark toxicity evaluation, cells were incubated with media 
containing photosensitizers (0-40 ?M) for 72 hours followed by MTT assay. The 
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photosensitizer efflux by ABC transporters was studies in MCF-7 and its multi-drug resistant 
sublines (MCF-7 TX400 and MCF-7 MX100) by adapting our protocol.  
7.2.7 Intracellular ROS Production using DCFDA 
Intracellular ROS production was examined via 2?,7??dichlorofluorescin 
diacetate (DCFDA; Thermo Fisher) assay. U87 cells were plated in 96-well black wall 
plates (Krystal) at a density of 2.2?104 cells/cm2. On the next day, cells were incubated 
with complete culture medium containing 0.25 ?M of NanoVP, free VP, or Liposomal 
VP for 24 hours. On day 3, prior to PDT, cells were washed twice with PBS and 
incubated with 10 ?M DCFDA for 30 minutes. Cells were irradiated with 690-nm red 
light (0-10 J/cm2, 50 mW/cm2; ML6600, Modulight). For the positive control, cells 
were incubated with 0.1 mL of 100 ?M H2O2 for 15 minutes. The fluorescence signal 
of the cleaved DCF probe (Excitation/Emission: 485/535?15 nm) was measured using 
a microplate reader (Synergy Neo2, BioTek). 
7.2.8 Mitochondrial Membrane Potential using TMRE 
Mitochondrial membrane potential (??m) was examined via TMRE (Abcam) 
assay. U87 cells were plated in 96-well black wall plates at a density of 2.2?104 
cells/cm2. After a 24-hour incubation with 0.25 ?M of NanoVP, free VP, or Liposomal 
VP, cells were washed twice with PBS and exposed to 690-nm light (0-10 J/cm2, 50 
mW/cm2). After PDT, cells were incubated with 250 nM TMRE for 25 minutes and 
washed twice with 0.2% bovine serum albumin within PBS. Samples were subjected 
to fluorescence readings (Excitation/Emission: 549/575?10 nm) using a microplate 
reader or fluorescence and phase contrast imaging (Lionheart, BioTek). Controls 
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include no light irradiation and incubation with 100 ?M of FCCP, a mitochondrial 
uncoupler, for 25 minutes. PDT and FCCP treated samples were normalized to a no 
treatment control. 
7.2.9 Caspase-3 Western Blots 
Expression of total and cleaved caspase-3 were examined using immunoblotting 
techniques. U87 cells were plated in 35-mm Petri dish (Falcon) at a density of 2.2?104 
cells/cm2. Cells were incubated with 0.25 ?M of NanoVP, free VP, or Liposomal VP for 24 
hours. Cells were washed twice with PBS and irradiated with 690-nm light (10 J/cm2, 50 
mW/cm2) in fresh culture medium. At 1 hour after PDT, cell lysates were collected in RIPA 
buffer supplemented with 1% protease inhibitor. Protein lysates (22 ?g) were separated on a 4-
12% precast Bis-Tris gel (NuPAGE) and transferred onto a nitrocellulose membrane. After 
blocking with Odyssey blocking buffer (Li-COR) for 1 hour, blots were incubated with primary 
antibody against beta-actin (#3700, cell signaling), caspase 3 (#9662, Cell Signaling,), or 
cleaved caspase 3 (#9661, Cell Signaling) overnight at 4 ?C. Blots were washed with 1X Tris-
Buffered Saline, 0.1% Tween? 20 Detergent buffer followed by incubation with IRDye 680RD 
goat anti-mouse secondary antibody (926-68070, Li-COR) or IRDye 800CW goat anti-rabbit 
secondary antibody (926-32211, Li-COR) for 1 hour at room temperature based on the 
supplier?s advice. Visualization of protein bands was developed using the LiCor ODYSSEY 
CLx (Li-COR). ?-actin serviced as a loading control. Band intensity was determined using 
LiCOR ODYSSEY CLx software. PDT treated samples were normalized to a no treatment 
control. 
7.2.10 Drug Efflux Assay 
To study ABC transporter-mediated photosensitizer efflux, MCF-7, MCF-7 
TX400, or MCF-7 MX100 were plated in 35 mm petri dishes (Falcon) at a density of 
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3.3 x104 cells/cm2. The next day, cells were incubated with 1 mM of NanoVP, free VP, 
or Liposomal VP for 4 hours at 37 ?C with or without ABC drug transporter inhibitors 
(ABCG2 inhibitor: 10 mM fumetrimorgin C; P-gp inhibitor: 2.5 mM tariquidar). 
Subsequently, cells were washed twice with PBS, trypsinized, and lysed in RIPA buffer 
at 4 ?C. VP fluorescence (Excitation/Emission: 435/650-750 nm) was measured using 
a multi-mode microplate reader and intracellular VP concentration was determined 
with appropriate VP standard curves. Intracellular VP was normalized to total cell 
protein. The total protein amount was determined using Pierce bichoninic acid protein 
assay (Thermo Fisher). 
7.2.11 In Vivo PDT and Photosensitizer Biodistribution 
Animal protocols were approved by University of Maryland, College Park 
Institutional Animal Care and Use Committee. Xenograft mouse model of GBM were 
established by subcutaneously injecting U87 cells (1?106 cells in PBS/Matrigel?) into 
the flank of J:NU mouse (4-5 weeks old, #007850, Jackson Laboratory). Tumor 
volumes were longitudinally monitored using calipers and calculated using the standard 
estimation formula, V=??length?width2, where length equals the maximum tumor 
diameter in millimeters and width equals the diameter that is perpendicular to the 
length. Treatments were initiated 2 weeks post-implantation when tumors reached ~100 
mm3. At 2 hours post-intravenous injection of photosensitizers (i.e., NanoVP or 
Liposomal VP; 0.25 mg/kg) or PBS, a vertical 690 nm laser beam (100 J/cm2, 100 
mW/cm2; ML6600, Modulight) was focused on the tumor to activate PDT. A cloth was 
used to protect animal skin from light exposure. Change in tumor volume was 
monitored for up to 2 months. The specific growth rate of tumors was estimated using 
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the equation (1/V)(dV/dt), where V is tumor volume and t is time. To examine 
photosensitizer biodistribution, tumor and normal tissues were collected at 2- and 24-
hours post-injection of photosensitizers. Fluorescence images of VP in organs, tumors, 
and standards containing known VP concentrations were acquired using a reflectance 
fluorescent microscope equipped with a 375 nm laser diode (6 mW, L375P70MLD, 
Thorlabs), a 692 nm filter (FF01-692/40-25 Semrock), and a 12-bit CCD camera (EM?
CCD, PCO, Kelheim, Lower Bavaria, Germany). Fluorescence images were processed 
by using a standard curve to convert the intensities into concentration using MATLAB. 
7.2.12 Photodynamic Opening of the Blood-brain Barrier in Rodents 
Animal protocols were approved by University of Maryland, School of 
Medicine Institutional Animal Care and Use Committee. Sprague-Dawley rats (4-5 
weeks old, Envigo) received intravenous injection of NanoVP (0.25 or 0.5 mg/kg) or 
5-aminolevulinic acid (5-ALA, 20 or 125 mg/kg) at 30 min or 4 hours before PDT. 
PDT parameters, including photosensitizer concentration, drug-light interval, 
irradiance and radiant exposure, were selected based our experience or other?s clinical 
work. At 30 minutes post-photosensitizer injection, rats were anesthetized with 
isoflurane, secured within a stereotaxic frame, and craniotomy was performed to 
expose the right cerebral hemisphere. PDT was performed by light activation of the 
exposed brain (NanoVP: 690 nm, 80 or 100 J/cm2; 5-ALA: 635-nm, 60 or 80 J/cm2; 40 
or 85 mW/cm2; ML6600, Modulight). At 90 minutes post-PDT, rats received 
intravenous injection of Evans blue dye (2%, 4 mL/kg) to determine the BBB integrity 
using imaging and extraction methods. In a small group of PDT animals that did not 
receive Evans blue injection, brain tissues were collected, sectioned, and processed for 
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(i) histological (H&E) analysis and (ii) Luxol Fast Blue staining of myelin/myelinated 
axons and Nissl bodies. H&E and immunohistochemistry slides were imaged using the 
whole slide Aperio AT2 scanner system (Leica Biosystems). All image analysis was 
accomplished using Aperio and Halo imaging analysis software (v3.3.2541.300; Indica 
Labs), and image annotations were performed by a pathologist (B.K). Fields were 
excluded if they contained large areas of artifact such as folds or tears. 
7.2.13 Rat Craniotomy 
Rats were anesthetized with isoflurane and secured within a stereotaxic frame. 
During the 30-minute drug-light interval, a craniotomy was performed. The top of the 
rat head was shaved, and the skin was removed. The periosteum was retracted to the 
edges of the skull. Using a micro drill (OmniDrill35, WPI), a circle about 4 mm in 
diameter was sketched into the right parietal bone. Drilling was stopped periodically, 
and PBS was applied to prevent heating. The drilled bone was removed and any 
bleeding that occurred from surgery was stopped prior to light irradiation. After a 30-
minute circulation period with Evans blue, rats were sacrificed via isoflurane overdose 
and decapitation, and brains were harvested. Digital images were immediately 
captured, and the brains were frozen at -80?C. 
7.2.14 Evans Blue Quantification 
Evans blue was quantified as previously described [498]. Briefly, the brain 
hemispheres were thawed on ice, minced, and incubated with formamide for 30 
minutes at 55?C followed by 4 days at room temperature. Samples were centrifuged to 
pellet the tissue and the optical density of Evans blue within the supernatant at 610 nm 
151 
 
 
was measured on a multi-mode microplate reader. Standard curves were used to 
determine the amount of Evans Blue in each brain hemisphere. 
7.2.15 Brain Histology 
The effects of PDT on healthy brain tissues were examined 2 hours after light 
irradiation. Brains were harvested and frozen within optimum cutting temperature 
compound (Tissue-Tek) at -80?C. Cryocut sections (5 ?m) from frozen rat brains were 
collected at the level of rostral and caudal to the treatment area and fixed in a mixture 
of glutaraldehyde and formaldehyde. H&E staining was performed using the Sakura? 
Tissue-Tek? Prisma? automated stainer.  The slides were hydrated and stained with 
commercial hematoxylin, clarifier, bluing reagent and eosin-Y. A regressive staining 
method was used.  This method intentionally overstains tissues and then uses a 
differentiation step (clarifier/bluing reagents) to remove excess stain. The slides were 
cover slipped using the Sakura? Tissue-Tek?Glass? automatic cover slipper and dried 
prior to review. For Luxol Fast Blue staining, sections were hydrated and stained 
overnight at room temperature.  Sections were immersed in 95% alcohol to remove 
excess stain. Slides were then washed in distilled water and quickly immersed in 0.05% 
lithium carbonate. Tissue sections were then placed in 70% ethanol until a sharp 
contrast between the corpus callosum and cortex developed. Sections were washed in 
distilled water, stain with cresyl violet acetate, differentiate in several changes of 95% 
alcohol, dehydrated with ethanol, cleared in xylene, and mounted. Slides were scanned 
at 20X using an Aperio AT2 scanner (Leica Biosystems) into whole slide digital 
images.  
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7.2.16 Statistical Analyses 
Results are presented in mean ? SEM. Statistical tests were carried out using GraphPad 
Prism (GraphPad Software). Specific tests and number of repeats are indicated in the figure 
captions. Reported P values are two-tailed. One-way ANOVA statistical tests and appropriate 
post hoc tests were carried out to avoid type-I error. 
7.3 Results  
7.3.1 Formulation and Characterization of NanoVP 
VP is barely soluble in water and prone to forming J-type aggregates in 
physiologically relevant buffers (Figure 7.1a). To date, intravenous delivery of VP in 
the clinic relies on the use of liposome, which serves as a solubilizer. Here, we 
successfully prepared a well-dispersed, carrier-free nanodrug of  VP (NanoVP) using 
the solvent-antisolvent precipitation technique (Figure 7.1b). After dissolving VP in 
100% DMSO at 7 mM, one part of sample was added dropwise to fifty parts of stirring 
water at room temperature and followed by dialysis against PBS at 4?C. Because 
DMSO and water are miscible, the introduction of VP DMSO solution in water creates 
local supersaturation of VP, which is thermodynamically unfavorable and could lead 
to the production of amorphous solids (precipitation) or crystals (crystallization). TEM 
micrographs revealed that NanoVP particles are spherical-like, amorphous, 
monodispersed, and ~100 nm in diameter (Figure 7.1c-e, Figure 7.2). The lack of 
electron diffraction further verified the amorphous structure of NanoVP (Figure 7.1f). 
TEM study using ionic liquid-treated samples showed that sample dehydration, 
destined for conventional TEM, has a negligible influence on the structure and size of 
NanoVP (Figure 7.3). Other attempts at synthesis, including adding VP (in 100% 
153 
 
 
DMSO) dropwise into PBS, resulted in uncontrollable agglomeration (Figure 7.4), 
potentially because the salts in PBS either served as nucleation sites or changed the 
charge of the dispersion [490]. 
Dynamic light scattering and laser Doppler electrophoresis measurements 
showed that NanoVP has a hydrodynamic diameter of 105.7?6.7 nm (PdI: 0.13?0.1) 
and a zeta potential of -32.9?0.3 mV (Figure 7.1g). Dialysis for purification and buffer 
exchange had minimal impact on NanoVP size (110.2?7.5 nm), PdI (0.13?0.2), and 
zeta potential (-29.6?6.7 mV) (Figure 7.1g). NanoVP has an entrapment efficacy of 
91.6?7.5%, reflecting minimal loss during sample preparation and dialysis. The 
loading capacity of NanoVP is 100% at 706,000?38,000 VP molecules per 
nanoparticle, which is approximately 1455-fold greater than Liposomal VP (485?75 
VP molecules per nanoparticle, Figure 7.5). The size of the NanoVP is tunable between 
65 nm and 150 nm by increasing the photosensitizer concentration in solvent (1-40 
mM, Figure 7.1h) or the DMSO:Water ratio (2-25%, Figure 7.1i). However, a starting 
VP concentration beyond 15 mM or a DMSO:Water ratio higher than 6% resulted in 
multi-peak size distribution (Figure 7.6a,b). NanoVP, synthesized using 1-7 mM VP 
and 2% DMSO:Water ratio, was found stable for over one year in water (pre-dialysis) 
and at least five months in PBS (post-dialysis) (Figure 7.7). Further studies indicated 
NanoVP stability is maintained via electrostatic repulsion forces between the particles 
(Figure 7.8). 
154 
 
 
 
Figure 8.1 Physical Assessment of the Amorphous Nanodrug of Verteporfin (NanoVP). 
(a) Representative digital image of free-form VP, Liposomal VP, and NanoVP in PBS. (b) 
Schematic depiction of the solvent-antisolvent precipitation method for preparation of 
NanoVP. Panel (c) depicts representative transmission electron microscopy (TEM) 
micrographs (Scale bar = 200 nm) and (d, e) close-up views of monodispersed NanoVP. Panel 
(f) shows a representative image of selected area electron diffraction of the amorphous 
NanoVP. (g) Representative intensity plots, the average hydrodynamic diameter, and the 
average zeta potential of NanoVP diameter pre and post dialysis. The hydrodynamic diameter 
and polydispersity index (PdI) of NanoVP (h) as a function of initial VP concentration in 
DMSO or (i) as a function of DMSO:water ratio. N ? 3, where N equals the number of samples.  
Error bars show the SEM. 
155 
 
 
Figure 8.2 Size Quantification of NanoVP TEM Micrograph Images. Black line shows the 
gaussian distribution curve of the data set. Bin size = 10 nm. n = 250 particles. 
 
 
Figure 8.3 Representative Ionic Liquid TEM Micrograph. A second electron imaging 
technique was used to verify the ?drying effect? did not impact NanoVP structure or 
morphology. Scale bar = 100 nm.  
 
 
Figure 8.4 Alternative NanoVP Synthesis Methods.  Representative digital images of (a) VP 
lyophilized powder was added to PBS and did not dissolve. Fully dissolved VP in DMSO was 
added into PBS (b) dropwise and (c) non-dropwise. Adding VP solution directly into PBS 
results in uncontrolled VP aggregation. (d) NanoVP synthesis following established 
preparticipation method in water. (e) NanoVP post dialysis in PBS. 
156 
 
 
 
 
Figure 8.5 Loading Capacity of NanoVP and Liposomal VP. (a) The amount of VP 
molecules per nanoparticle was determined using the following formula: VPparticle = 
Concentrationsolution * NA / particles per ml solution. (b) Particles per ml solution. Two tail t-
test was used to calculate significant differences, where **** p < 0.0001. N ? 3.  Box plots 
show the mean and the likely range of variation. Error bars show the maximum and minimum 
values.  
 
 
Figure 8.6 NanoVP Diameter. Representative intensity plots of NanoVP as a function of (a) 
DMSO: Water ratio and (b) initial VP concentration. 
 
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Figure 8.7 NanoVP Shelf Life. (a) NanoVP diameter and PdI in PBS and (b) representative 
intensity plots in water. N?3, where N equals the number of samples.  
 
Figure 8.8 Probing NanoVP Electrostatic Stabilization. Representative digitals images of 
NanoVP coated polymers. NanoVP was synthesized with (left) neutral polymer poloxamer 407 
or (right) cationic polymer polyethylenimine. Poloxamer 407 had no impact on NanoVP 
stability, size, nor PdI. Polyethylenimine caused NanoVP to form large aggregates.  N?3, where 
N equals the number of samples. 
7.3.2 Photo-activable NanoVP 
VP is a modified porphyrin derivative that displays a chlorin-type spectrum in 
organic solvents. In DMSO, the absorption spectrum of VP is characterized by a 
distinct Q band in the near-infrared region at 687 nm and a strong Soret band at 435 
nm (Figure 7.9a). The absorption spectra of NanoVP, free-form VP, and Liposomal 
VP were identical in DMSO, where VP is mainly in its monomeric form. At an 
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excitation wavelength of 435 nm, the fluorescence emission spectra of NanoVP in 
DMSO can be recorded around 700 nm (Figure 7.9a). When NanoVP is well-dispersed 
in PBS, a broadening of the Soret band and a red-shifted Q band were observed (Figure 
7.9b). This suggests that monodispersed NanoVP consists of self-assembled J-
aggregates. On the contrary, liposomes maintain some monomeric form of VP in PBS, 
showing a minimal decrease in the Soret band and without a red-shift displacement of 
the Q band as expected (Figure 7.9b). Driven by the spatial confinement of high-
number VP in a 100 nm diameter particle, NanoVP generates extensive fluorescence 
self-quenching up to 328-fold in PBS in a concentration-dependent manner (Figure 
7.9c). The degree of self-quenching in free-form VP or Liposomal VP were only up to 
78-fold or 2-fold, respectively. VP quenching can decrease the photochemical 
production of singlet oxygen (1O2) (a Type II reaction). Upon light activation (690 nm, 
10 J/cm2, 10 mW/cm2), highly quenched NanoVP and free-form VP in PBS did not 
produce any significant amount of 1O2, as indicated by the minimal SOSG fluorescence 
signal (Figure 7.9d). On the contrary, light activation of Liposomal VP generates an 
up to 34-fold higher SOSG signal, compared to NanoVP. Similar to that observed in 
clinical PDT practice, we also confirmed that light activation of NanoVP, free-form 
VP, or Liposomal VP results in limited photothermal effects (?T=2-3?C; Figure 7.10).  
The highly quenched NanoVP can be dequenched in the presence of serum 
proteins and in cancer cells for photosensitized 1O2 production. Figure 7.9e shows that 
increasing the serum protein level from 0.15 to 10 % v/v decreases NanoVP quenching 
from 92-fold to 3.2-fold. Disaggregation of NanoVP in serum-containing media and 
U87 cells was further studied by longitudinal monitoring the VP fluorescence signal 
159 
 
 
(Figure 7.11). After adding NanoVP to 10% v/v serum-containing media, a rapid 
increase in VP fluorescence within 30 minutes is followed by the plateau of the signal, 
whereupon the signal remains constant at 38% of that fully dissolved in DMSO (Figure 
7.11a). Similar trends were observed after adding NanoVP to U87 cells in serum-
containing media (Figure 7.11b) or PBS (Figure 7.11c), where fluorescence recovery 
was the highest in NanoVP samples, compared to free-form VP and Liposomal VP. To 
check the recovery of photochemical activity of NanoVP in U87 cancer cells, DCFDA 
was used as a fluorescent probe for the detection of ROS generation (Figure 7.9f). 
Upon light activation (690 nm, 10 J/cm2, 50 mW/cm2), disaggregated NanoVP 
produced significantly higher intracellular ROS (~2-fold, P < 0.001), compared to that 
of Liposomal VP and the positive control (100 ?M H2O2). 
160 
 
 
 
Figure 8.9 Photochemical Characterization of NanoVP, Free-form VP, and Liposomal 
VP. (a) Representative absorbance and fluorescence (FL) spectra of 5 ?M NanoVP, free-form 
VP, and Liposomal VP in DMSO. VP was excited at excitation 435 ? 10 nm. (b) Representative 
absorbance spectra of NanoVP, free-form VP, and Liposomal VP in PBS (red) and DMSO 
(blue). (c) Fluorescence quenching, the maximum fluorescence intensity of 0.625-5 ?M 
NanoVP, free-form VP, and Liposomal VP in DMSO (FLDMSO) divided by the maximum 
intensity within PBS (FL0). (d) Singlet oxygen production from NanoVP, free-form VP, and 
Liposomal VP-mediated PDT (10 J/cm2, 10 mW/cm2). (e) Fluorescence quenching of NanoVP 
as a function of serum concentration in PBS. (f) The intracellular ROS production from 
NanoVP, free-form VP, and Liposomal VP-mediated PDT (10 J/cm2, 50 mW/cm2). One-way 
ANOVA with multiple comparison test was used to calculate significant differences, where 
*p<0.05, ** p < 0.01, *** p < 0.001. N ? 3, where N equals the number of wells conducted 
over 3 trials.  Error bars show SEM. 
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Figure 8.10 High Power Light Activation of VP Results in Limited Photothermal Effects. 
NanoVP, free-form VP, or Liposomal VP (40 ?M) in PBS were light activated (50 J/cm2, 0.5 
W/cm2) and solution temperature was measuring. Error bars show SEM.  N?6, where N equals 
the number of wells conducted over 3 trials. 
 
Figure 8.11 NanoVP Fluorescence 
Recovers in Physiologically Relevant 
Mediums. (a) The time-dependent 
fluorescence signal (Excitation/Emission: 
435/700) of 5 ?M in PBS, DMSO, and 
complete cell culture medium supplemented 
with 10% FBS at 37?C. (b) The time-
dependent normalized fluorescence signal of 
NanoVP, free-form VP, and Liposomal VP 
when incubated with a monolayer of U87 
cells within cell culture medium or (c) PBS 
over a 3-hour period at 37?C. N?6, where N 
equals the number of wells conducted over 3 
trials. 
 
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7.3.3 In vitro NanoVP Uptake and PDT Efficacy 
The therapeutic efficacy of PDT depends on the concentration of 
photosensitizer in cancer cells. U87 cells treated with NanoVP resulted in ~2-fold 
higher intracellular concentration of VP (129.8?29.1 ng per mg of protein) compared 
to using Liposomal VP (66.2?8.1 ng per mg of protein) (Figure 7.12a). The uptake of 
NanoVP in U87 cells was similar to free-form VP. MTT assay revealed that NanoVP 
has a stronger anti-cancer PDT effect than Liposomal VP (Figure 7.12b). NanoVP and 
free-form VP exhibited the lowest half-maximal inhibition concentration (IC50) of 0.20 
?M?J/cm2 in U87 cells, while Liposomal VP displayed a 4-fold higher IC50 of 0.81 
?M?J/cm2. The cellular uptake and PDT efficacy were further examined in non-
cancerous 3T3 mouse fibroblast cells (Figure 7.13). The intracellular uptake of 
NanoVP in 3T3 cells was 49.9% greater than Liposomal VP. This resulted in a 2-fold 
IC50 increase from ~0.4 ?M?J/cm
2 for NanoVP to 0.8 ?M?J/cm2 for Liposomal VP. 
NanoVP-PDT in U87 cells has a 50% lower IC50 compared to 3T3 cells, while the IC50 
of Liposomal VP-PDT remained similar in both U87 cancer 3T3 cells. These results 
indicate that the intracellular uptake and anti-cancer PDT efficacy of NanoVP is 
significantly higher than that of Liposomal VP.  
We and others have previously shown that VP preferentially accumulates in 
mitochondria and light activation of VP depolarizes the mitochondrial membrane 
potential (??m) to trigger apoptosis [21, 499, 500]. Here, we reported that light 
activation (690 nm, 10 J/cm2, 50 mW/cm2) of NanoVP results in the largest degree of 
??m depolarization, as indicated by TMRE assay (Figure 7.12c,d). Quantification and 
normalization of the TMRE signal reveals NanoVP-PDT depolarizes ??m by nearly 
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60%, while Liposomal VP-mediated PDT results in a modest depolarization of ??m 
by ~35%. The potent mitochondrial oxidative phosphorylation uncoupler, FCCP, was 
used as a positive control. PDT-induced mitochondria damage can usually trigger the 
intrinsic apoptosis pathway leading to caspase-3 activation [501] within a few hours 
after light activation [502-504]. Immunoblotting results indicate that PDT using 
NanoVP or free-form VP induced a ~2.4-fold increase in cleaved caspase-3 formation 
one hour post light activation (Figure 7.12e,f). In addition, NanoVP and free-form VP-
mediated PDT significantly reduced total (pro) caspase-3 expression (i.e., non-
activated caspase-3) by ~50%, resulting in a cleaved-to-total caspase 3 ratio of ~4.4. 
Liposomal VP-PDT resulted in the lowest ratio of ~2.2. Neutral red uptake assay 
revealed that light-activation (690 nm, 1 J/cm2, 50 mW/cm2) of NanoVP, free-form VP, 
or Liposomal VP does not affect lysosomal integrity in U87 cancer cells. (Figure 7.14). 
  We have previously shown that free-form VP can be effluxed by ABC drug 
transporters, ABCG2 and P-gp, in cancer cells and thereby reducing PDT efficacy 
[196]. Using a human breast cancer cell line MCF-7, and their sub-lines (ABCG2 MCF-
7 MX100 and or P-gp-overexpressing MCF-7 TX400) [196, 505], we discovered that 
NanoVP is a substrate of ABCG2 and P-gp (Figure 7.15). Our studies further 
demonstrated the use of fumitremorgin C (ABCG2 inhibitor) and tariquidar (P-gp 
inhibitor), can improve the accumulation of NanoVP in MCF-7 MX100 and MCF-7 
TX400 cells, respectively. 
In general, PDT requires very low doses of photosensitizers (nM-?M) to be 
effective. Recent clinical and preclinical studies have reported that high doses of VP 
(mM) can induce potent ?dark? toxicity (without light activation) in cancer cells, 
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including GBM (NCT04590664) [506-508]. In complete darkness, NanoVP exhibited 
an IC50 of 12-14 ?M in U87 cells and 3T3 cells, similar to that of free-form VP (Figure 
3g, Figure 7.16). NanoVP treatment at, or above, 20 ?M significantly reduced cell 
viability by 75%, while Liposomal VP exhibited no significant cytotoxicity presumably 
due to the poor intracellular VP uptake (Figure 7.12h,i). 
165 
 
 
 
Figure 8.12 In Vitro PDT Efficacy of NanoVP within U87 Cells. (a) The intracellular VP 
concentration determined via extraction method at 24 hours post incubation with 0.25 ?M 
NanoVP, free-form VP, or Liposomal VP. (b) Cell viability measured via MTT assay 24 hours 
after NanoVP, free-form VP, or Liposomal VP-mediated PDT (0-5 J/cm2, 10 mW/cm2). (c) 
Representative fluorescence images of the TMRE probe and phase constant images of U87 
cells at 30 minutes after PDT. (d) Mitochondrial membrane depolarization quantified via 
TMRE probe 1-hour after NanoVP, free-form VP, or Liposomal VP-mediated PDT (10 J/cm2, 
166 
 
 
50 mW/cm2). Depolarization is calculated using the following formula: ??m = 1-(Tf/Tc), 
where Ta is the TMRE fluoresce signal from FCCP or a treatment group, and Tc is the TMRE 
fluoresce signal from the control group. (e) Representative immunoblotting showing changes 
in total and cleaved caspase 3 expression in U87 cells at 1 hour post PDT. (f) Quantification of 
total and cleaved caspase 3 expression in U87 cells at 1 hour post PDT (10 J/cm2, 50 mW/cm2). 
A two-tail (total) and one-tail (cleaved) t-test was used to calculate significant differences. (g) 
Cell viability measured via MTT assay 72 hours after incubation with non-PDT activated (0 
J/cm2, 0 mW/cm2) NanoVP, free-form VP, or Liposomal VP. (h) Representative fluorescence 
images of NanoVP, free-form VP, or Liposomal VP within U87 cells. Scale bar = 50 ?m. (i) 
Quantitative analyses of NanoVP, free-form VP, or Liposomal VP fluorescence signal in U87 
cells. Fluorescence signals were normalized to the largest signal for each concentration. One-
way ANOVA with multiple comparison test was used to calculate significant differences, 
where * p < 0.05, ** p < 0.01, and *** p < 0.001. N?6, where N equals the number of wells 
conducted over 3 trials.  Error bars show SEM. 
 
 
 
Figure 8.13 PDT Efficacy within 3T3 Fibroblast Cells.  (a) The intracellular VP 
concentration determined via extraction methods at 24 hours post incubation with 0.25 ?M of 
NanoVP, free-form VP, or Liposomal VP. (b) Cell viability measured via MTT assay 24 hours 
after NanoVP, free-form VP, or Liposomal VP-mediated PDT (0-5 J/cm2, 10 mW/cm2). One-
way ANOVA with multiple comparison test was used to calculate significant differences, 
where *** p < 0.001. Error bars show SEM. N?6, where N equals the number of wells 
conducted over 3 trials.  
 
 
 
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Figure 8.14 VP-mediated PDT does not Induce Lysosomal Damage. Lysosomal damage 
assessed via neutral red uptake assay (a) 1 and (b) 24 hours post PDT (1 J/cm2, 50 mW/cm2) 
of 0.25 ?M NanoVP, free-form VP, or Liposomal VP. One-way ANOVA with multiple 
comparison test was used to calculate significant differences, where * p < 0.05, N?3, where N 
equals the number of wells conducted over 3 trials.  Error bars show SEM.  
Figure 8.15 NanoVP is a Substrate for ABC Transporters ABCG2 and P-gp. Intracellular 
VP concentration determined via extraction methods at 4 hours post incubation with 1 ?M 
NanoVP or free VP. Breast cancer cells overexpressing (a) P-gp (MCF-7 TX400) or (b) 
ABCG2 (MCF-7 MX100), and parental MCF-7 cells were incubated with and without 
inhibitors (10 ?M fumitremorgin C for ABCG2 or 2.5 mM tariquidar for P-gp). Two-way 
ANOVA with multiple comparison test was used to calculate significant differences, where * 
p < 0.05, ** p < 0.01, N?3, where N equals the number of wells conducted over 3 trials. Error 
bars show SEM.  
Figure 8.16 NanoVP Dark Toxicity in 3T3 Cells. Cell viability measured via MTT assay 72 
hours after incubation with non-PDT activated (0 J/cm2, 0 mW/cm2) NanoVP, free-form VP, 
or Liposomal VP. N?6, where N equals the number of wells conducted over 3 trials.  
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7.3.4 NanoVP Enhances Anti-Glioblastoma PDT 
To assess the efficacy of NanoVP-PDT in controlling GBM tumors in vivo, 
PDT was performed fourteen days following subcutaneous implantation of U87 cells 
in mice, when tumors reached approximately 100 mm3 in volume (Figure 7.17). 
Liposome VP-PDT served as the clinical gold standard. For PDT, light (690 nm) 
irradiation was performed two hours after a single intravenous injection of NanoVP or 
Liposomal VP at 0.5 mg/kg. At 2-11 days after treatment, NanoVP-PDT reduced tumor 
volume by up to 35%, whereas continued tumor growth was observed in ?no-treatment? 
control and ?Liposome VP-PDT? over this same period (Figure 7.17a). At 21 days 
post-treatment, the mean tumor volume reduction in mice treated with ?NanoVP-PDT? 
and ?Liposome VP-PDT? was 70% and 44%, respectively, compared to ?no-treatment? 
control animals. These results suggest that NanoVP is a superior anti-tumor PDT agent 
compared to Liposomal VP. Further analysis revealed that animals treated with 
?NanoVP-PDT? experienced a significant tumor growth inhibition at a specific growth 
rate of around -4.3 %/day between days 0-11 after treatment (Figure 7.17b). With only 
a single cycle of treatment, tumor regrowth can be observed 11 days after ?NanoVP-
PDT? with a specific growth rate of approximately 5.3 %/day, similar to that of ?no-
treatment? and ?Liposome VP-PDT?. In ?NanoVP-PDT? and ?Liposome VP-PDT? 
treated animals, the change in mouse weight was consistent with ?no-treatment? mice, 
indicating that PDT does not appreciably add to the systemic toxicity (Figure 7.18). It 
is noteworthy that with a single cycle of treatment, only modest survival benefits were 
observed after ?NanoVP-PDT? (median survival: 33 days) and ?Liposome VP-PDT? 
(median survival: 31 days), compared to ?no-treatment? control (median survival: 25) 
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(Figure 7.17c). These results point out challenges in achieving meaningful 
improvements in treatment response for GBM and highlight the need for combination 
strategies developed to provide durable tumoricidal control. In addition to changes in 
tumor volume, noticeable visual differences on tumor surfaces were observed (Figure 
7.17d). At six days post-treatment, ?NanoVP-PDT? resulted in a significantly larger 
dark, possibly necrotic, area that covered the majority of the tumor (30.5 mm2), while 
?Liposomal VP-PDT? led to a smaller (13.6 mm2) and more localized necrotic patch 
(Figure 7.17d,e). Finally, fluorescence imaging of VP in mouse tissues revealed that 
NanoVP and Liposomal VP have a similar biodistribution profile at 2 and 24 hours post 
intravenous injection (Figure 7.17f, Figure 7.19). 
 
Figure 8.17 The Phototoxicity and Biodistribution of NanoVP and Liposomal VP in an 
U87 Glioblastoma Xenograft Mouse Model. PDT treatment (100 J/cm2, 100 mW/cm2) was 
initiated ~14 days after subcutaneous U87 cancer cell implantation when tumor volumes 
reached approximately 100 mm3. Mice were randomized into groups that received (i) no-
treatment, (ii) Liposomal VP 0.5 mg/kg, (iii) NanoVP 0.5 mg/kg. (a) Tumor volume was 
longitudinally monitored and calculated using the standard estimation formula, 
V=??length?width2, where length equals the maximum tumor diameter in millimeters and 
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width equals the diameter that is perpendicular to the length. Tumor volumes were normalized 
to the initial volume at the time of treatment. (b) The specific growth rate of tumors 0-11 
days and 11-25 days post PDT were determined using the following formula: specific growth 
rate = (1/V)(dV/dt), where V is tumor volume and t is time. (c) Kaplan-Meier plot of tumor 
diameter greater than 1.5 cm (n = 7?8 animals per group). (d) Quantification of the surface area 
above the tumor that was impacted by PDT treatment. (e) Representative digital images of 
tumors 6 days post PDT treatment. (f) Quantitative analyses of VP fluorescence intensity within 
the tumor and organs, 2 and 24 hours after VP injection. One-way ANOVA with multiple 
comparison test was used to calculate significant differences, where * p < 0.05, ** p < 0.01, 
and *** p < 0.001. N?7, where N equals the number of animals conducted over 2 trials. Error 
bars show SEM. 
Figure 8.18 Mouse Weight post PDT Treatment. PDT had no impact on weight. N?5. 
 
Figure 8.19 VP Biodistribution in Mice Organs and U87 Tumors. Representative 
fluorescence images of NanoVP and Liposomal VP in mice organs and U87 tumors at both 2- 
and 24-hrs post i.v. injection. Scale bar = 3 mm.   
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7.3.5 NanoVP-PDT Modulation of the Blood-brain Barrier in Rats 
The BBB remains a critical obstacle to the effective treatment of GBM and 
other central nervous system diseases. Low-dose PDT priming, an enabling technology 
for minimally invasive tumor treatments, allows transient opening of the BBB through 
modulating endothelial cell-cell junction phenotype. Here, we evaluate the utility and 
safety of low-dose NanoVP-PDT priming for spatially targeted BBB opening in 
healthy rats with intact BBB (Figure 7.20a). The 5-ALA (Gliolan?), an FDA-
approved photosensitizer prodrug routinely used for PDT opening of the BBB, served 
as benchmark group is our study. We identified a 5.5-fold increase of total Evan Blue 
accumulation in NanoVP-PDT-treated brain compared to 5-ALA-PDT treated brain 
tissues (P < 0.05) (Figure 7.20b,c). The spatiotemporal selectivity of PDT confines 
Evan Blue delivery to the right-brain hemisphere where the light is directed, thereby 
reducing normal tissue damage. The use of the longer activation wavelength 
photosensitizer, NanoVP (690 nm), for deeper tissue penetration is also an advantage 
over using the 5-ALA-induced protoporphyrin IX (PpIX, 635 nm) (Figure 7.20d). We 
showed that 690 nm light activation of NanoVP improves Evan Blue delivery at further 
depths in rat brains (visible up to 4-5 mm), compared to 635 nm light activation of 5-
ALA-induced PpIX (visible up to 1 mm). Histological changes of brain tissue after 
PDT were studied microscopically on sections stained with H&E or Luxol Fast blue. 
Compared to the left-brain hemisphere that did not receive treatment due to the intact 
skull, the right-brain hemisphere did not have any signs of low-dose PDT priming-
induced damage. H&E staining revealed no detectable lesions at the site of treatment 
(right hemisphere) and, Luxol fast blue staining was symmetrical with no evidence of 
172 
 
 
demyelination (Figure 7.20e). Alternatively, rats that received traditional high dose 
PDT had signs of brain damage within the neuropil parenchyma (brain cortex) (Figure 
7.21). This result agrees with previous studies that show dose escalation results in 
increased signs of edema and brain damage [17]. 
 
Figure 8.20 PDT-induced Blood-brain Barrier Opening within the Rat Brain. (a) 
Schematic depiction of experimental design. NanoVP (0.25 mg/kg) or 5-aminolevulinic acid 
(5-ALA, 20 mg/kg) were intravenously administered 30 minutes PDT (NanoVP: 690 nm, 80 
J/cm2; 5-ALA: 635 nm, 80 J/cm2; 85 mW/cm2) was performed on the exposed right brain 
hemisphere. After 90 minutes, Evans blue was intravenously administered to the rats and 
circulated for 30 minutes prior to brain harvesting. (b) Representative top and cross-sectional 
images of Evans blue within the brain after PDT-induced BBB opening (c) Quantification of 
Evans blue extracted from the right brain hemispheres (n ? 3 rats per group, background 
subtracted). (d) Quantification of maximum depth that extravasated Evans blue can be 
visualized within the brain. (e) Representative histopathology of rat brain tissue after traditional 
and low-dose PDT treatment. Photomicrographs of brain sections stained with H&E and Luxol 
173 
 
 
fast blue. Two tail t-test was used to calculate significant differences, where n.s. indicates not 
significant (p > 0.05), * p < 0.05. Error bars show SEM. N ? 3. 
 
 
Figure 8.21 Traditional PDT Damages Healthy Brain Tissue. Representative 
photomicrographs of brain sections stained with H&E and Luxol fast blue. Photosensitizers 
(0.5 mg/kg VP or 125 mg/kg 5-ALA) were photoactivated (VP: 100 J/cm2, 40 mW/cm2; 5-
ALA: 60 J/cm2, 40 mW/cm2) 30-minutes (for VP) or 4-hours (for 5-ALA) after intravenous 
injection. Brains were harvested 90 minutes post PDT. N?3. 
7.4 Discussion 
PDT using Liposomal VP has already helped hundreds of thousands of patients 
globally with wet age-related macular degeneration [145]. Since its FDA approval in 
2001, Liposomal VP has been tested clinically to treat a wide range of cancers, 
including primary breast, retinoblastoma, and refractory brain tumors. Liposomal VP-
PDT is currently being evaluated for unresectable solid pancreatic tumors or advanced 
pancreatic cancer (NCT03033225), and the chemotherapeutic effects of Liposomal VP 
are being evaluated in patients with recurrent high-grade EGFR-mutated glioblastoma 
(NCT04590664). Free-form VP is not used clinically because the large VP 
agglomerations created in aqueous buffers will negatively impact its pharmacokinetics 
and singlet oxygen yield. Pure-drug nanoparticle delivery systems offer tools to 
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improve the pharmacokinetic profile of hydrophobic drugs and minimize the reliance 
on solubilizing agents [509]. In this study, we report a new pure-drug nanoparticle of 
VP (NanoVP) that significantly improves photosensitizer delivery to cancer cells. Our 
in vivo findings and results show the safety, feasibility, and potential utility of NanoVP 
for PDT of gliomas, as well as BBB opening to enhance drug delivery with no evidence 
of microscopic normal tissue injury. 
Solvent antisolvent precipitation is a simple and reproducible formulation 
strategy to synthesize stable nanoparticles of VP. The size of NanoVP is tunable 
between 65 nm and 150 nm by increasing the photosensitizer concentration in solvent 
or the DMSO:Water ratio. Similarly, others demonstrated a larger initial concentration 
and higher solvent:antisolvent ratio could increase the size of nanodrug paclitaxel 
[510]. Synthesis optimization revealed that monodispersed NanoVP can be produced 
using an initial VP concentration of less than 15 mM and a DMSO:Water ratio below 
6%. A future direction includes examining the impact of solvent type (e.g., ethanol or 
methanol) on NanoVP formation, size, and stability. The amorphous structure of the 
prepared NanoVP, showed by X-ray diffraction data, represents another major 
advantage in the quest to immediately convert quenched NanoVP into its un-quenched 
molecular form in presence of the serum components or cells, which recovers 
fluorescence signal and PDT capability.  
Previous studies have also shown that pure-drug nanoparticles can improve 
cellular pharmacokinetics while liposomes hinder therapeutic uptake [489, 511, 512]. 
As a direct result of improved cellular accumulation, NanoVP was found to be a more 
potent PDT and chemotherapeutic agent than the traditional Liposomal VP. This result 
175 
 
 
is similar to previous studies that have shown free-form drugs, including doxorubicin 
[513], and paclitaxel [511], are uptaken more effectively in a variety of cancer cells 
compared to their liposomal counterparts. Free-form VP is lipophilic and can passively 
diffuse through the cell membrane [514, 515]. It is speculated that NanoVP had a 
similar accumulation rate as free-form VP, as NanoVP is readily dissociated into its 
molecular form in presence of the serum components or cells. When VP is encapsulated 
within a liposome, the mode of cellular entry is generally altered from passive diffusion 
to endocytosis [516], hindering the overall drug accumulation rate. We further present 
evidence that NanoVP-PDT, similar to free-form VP-PDT, effectively produces 
intracellular ROS, induces mitochondrial membrane potential (??m) depolarization, 
and initiates intrinsic apoptosis upon light activation. Typically, PDT requires very low 
doses of photosensitizers (in the nanomolar to micromolar range) to be effective. 
Recent clinical and preclinical studies have reported high concentrations of VP (in the 
millimolar range) can induce ?dark? cytotoxicity in cancer cells, including GBM 
(NCT04590664) [506-508]. A 10-100-fold higher dose of VP, compared to the PDT 
dose, has been shown to downregulate Bcl-2, disrupt the YAP/TAZ-TEAD complex 
interaction, and induce cancer cell death without light activation [506, 507]. We 
demonstrated that the enhanced cellular uptake of NanoVP resulted in superior non-
PDT killing effects. Alternatively, Liposomal VP alone did not inhibit cancer growth 
at the same millimolar incubation concentrations.  
While this report offers many findings and formulation advances related to VP-
PDT, there are some notable limitations. In subcutaneous glioma mouse models, 
NanoVP-PDT effectively reduced initial tumor growth, outperforming Liposomal VP-
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PDT. Although VP-PDT has been used successfully in the clinic to treat different 
cancers, a single cycle of PDT, like most cancer regimens, is insufficient to achieve a 
long-term survival. In our study, NanoVP increased the median survival of mice 
bearing subcutaneous tumors by over a week. This result is similar to our previous 
study, where we demonstrated a single dose of VP-PDT (0.25 mg/kg) could reduce 
OVCAR-5 tumor volume by up to 55% but did not increase overall survival [278]. 
Similar to chemotherapy, radiation and immunotherapy, NanoVP-PDT would require 
multiple treatment cycles to effectively eradicate the entire cancerous cell population 
and warrants further investigation. Nonetheless, a single cycle of NanoVP-PDT 
remains an attractive adjuvant therapy that can be combined with surgery. In a prior 
clinical study, Liposomal VP-PDT decreasing tumor volume could convert patients 
with previously unresectable pancreatic tumors to become candidates for surgery [168]. 
In addition surgery, PDT has been successfully combined with chemotherapy, radiation 
or immunotherapy, due to different mechanisms of action and their non-overlapping 
side effects [19]. Biodistribution analysis revealed NanoVP tended to have a higher 
tumor accumulation, while Liposomal VP accumulation more within the liver, 
pancreas, and spleen. Taken together, these results suggest that NanoVP could be an 
adequate replacement for our pegylated Liposomal VP formulation. A further 
examination of the pharmacokinetics and PDT parameters (e.g., light fluence, drug-
light interval) will be important next steps in determining if NanoVP is a good 
replacement for the clinically used non-pegylated liposome formulation.  
Drug delivery to brain tumors remains especially difficult due to the BBB, a 
specialized network of capillaries in the brain. The high degree of tightness and 
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integrity imposed by the BBB limits the effective delivery of more than 98% of small-
molecule therapeutics to the brain [7]. While many treatment modalities have shown 
promise in opening (i.e., permeabilizing) the BBB and enhancing brain drug delivery, 
they often result in edema and neurotoxicity, limiting their clinical use [8-11, 117]. 
Numerous groups have shown the potential of light-activated 5-ALA (Gliolan?)-
induced protoporphyrin IX (PpIX) to open the BBB in rodents [16, 517]. Hirschberg 
and colleagues showed larger doses of 5-ALA-PDT could open the BBB for up to 72 
hours [14, 15], but treatment resulted in early signs of necrosis up to 5 mm away from 
the primary brain tumor. In addition, non-tumor-bearing animals that received 125 
mg/kg 5-ALA and 54 J experienced a 50% mortality within 5 days of treatment. Here, 
we studied the effects of NanoVP-PDT at subtherapeutic doses to achieve 
photodynamic priming [175, 518]. This approach is particularly relevant for brain 
tumors, where surrounding healthy tissue damage and intratumoral drug accumulation 
are a major concern. In this study, we demonstrated that NanoVP-PDT mediates BBB 
opening to increase in EB accumulation in the brain by 5.5-fold compared to using 5-
ALA-PDT. More importantly, we show for the first time that low-dose PDT using 
NanoVP did not result in healthy brain tissue damage.  
Finally, we believe that intraoperative NanoVP-PDT has a high translational 
potential. Surgery followed by PDT of GBM has been studied in the clinic (e.g., 
NCT03048240, NCT03897491, NCT00003788) [143, 169]. Despite promising results, 
the side effects of PDT remain a major concern [14, 16, 519-521]. We believe the most 
advantageous, safe initial application of low dose PDT is following tumor resection 
while patients are in the operating room. In this context, patients will receive PDT 
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intraoperatively via fiber optic light conduits placed within the resection cavity [143, 
169]. Our ultimate goal is to open the BBB and sterilize unresectable, brain-invading 
tumor cells in real-time, under controlled conditions, during open surgery. Light 
penetration depth (defined as the depth at which the incident optical energy drops to 
37%, 1/e) is typically 1-4 mm in most tissues [125]. Others and we have shown that 
diffused light (i.e., the remaining 37% optical energy) could reach up to 1.5-2 cm in 
tissues [185-187]. Wilson, Perria, and Cheng have demonstrated that light (635 nm) 
activation of porphyrin induces tumor killing up to 1.5 cm deep in the brain [185-187]. 
In rat brain tissues, we have shown that diffused light (690 nm) can reach up to 1.5-2 
cm deep in rat brains to activate VP for effective PDT [22]. This suggests that VP-PDT 
is sufficient to manage post-surgical residual GBM cells (typically within 1-2 cm from 
the border of the original lesion) that are responsible for ~80% recurrence. Moreover, 
the self-limiting depth of effect avoids non-specific priming of the underlying tissue. 
Rarely, the GBM recurrence occurs distant beyond 2 cm of the resection cavity [522]. 
For distant recurrence (?ghost? cells) or inoperable GBM, light can be delivered to the 
entire tumor using stereotactically placed fibers in the clinic [80, 256]. Our next step is 
to investigate NanoVP-PDT after primary tumor resection to open the BBB and 
enhance the subsequent delivery of chemotherapy or immunotherapy to invasive, 
microscopic cancer cells embedded within healthy brain tissues. 
7.5 Conclusions 
The BBB remains a critical obstacle to the effective treatment of central nervous 
system diseases, such as GBM?the most common and deadliest form of primary brain 
cancer in adults. In this study, we report a novel, amorphous, pure-drug nanoparticle of 
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verteporfin for PDT of GBM and controlled BBB opening in rats. The findings revealed 
the capability of this new verteporfin nanoformulation to provide a safer and more 
effective option for cancer PDT and brain drug delivery applications. Our findings 
demonstrate the capability of this nanoformulation to control the opening of BBB with 
spatiotemporal precision and no sign of normal tissue injury, providing opportunities 
for improved delivery of chemotherapies, immunotherapies, nanomedicines to brain 
tumors. 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
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9. Chapter 8. Development of a High-Throughput Screening 
Strategy for Discovery of Human and Zebrafish ABC 
Transporter Substrates 
 
8.1 Introduction 
The human brain, one of the most vascularized organs in the body, is protected 
from circulating toxins (and potential therapeutics) that might gain entry by specialized 
capillaries known as the BBB [83]. The BBB is part of a larger and more complex 
neurovascular unit, composed of endothelial cells, neural cells (i.e., astrocytes, 
neurons), pericytes, microglia, and the basement membrane. It regulates nutrient 
transport, maintains brain homeostasis, and protects the central nervous system [81]. 
The BBB is composed of a physical and active barrier between the circulatory system 
and the brain parenchyma. Tight junction transmembrane complexes located on the 
apical region of the intracellular cleft prevent the diffusion of charged, large (>400 Da), 
and hydrophilic (>?8 hydrogen bonds) molecules between the apical and basal sides of 
the cell [7, 82]. The endothelial cells are also lined with ABC transporters that redirect 
toxins and therapeutics back into the bloodstream [110]. ABC transporters, localized 
to the luminal surface of endothelial cells, transport a wide range of substances against 
their concentration gradient via ATP hydrolysis [110]. Two of the highly expressed 
ABC transporters at the human BBB are Pg-p (also known as ABCB1) and ABCG2. 
The BBB is a major hurdle when delivering therapeutics to the brain [88]. Decades of 
research has shown the difficulties in treating brain malignancies as many therapeutics 
cannot successfully enter the brain because they are transported by one or both ABC 
transporters [523]. 
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Mouse models have consistently demonstrated the major role ABC transporters 
play in keeping chemotherapeutic agents out of the brain.  One of the first murine 
studies demonstrated deletion of the Abcb1a gene resulted in a 20-fold increase in brain 
vincristine levels compared to wild-type mice [524]. Upon identification of ABCG2 as 
a major component of the BBB, studies examined the effects of deleting Abcb1a and 
Abcb1b (the two murine homologs of human ABCB1), Abcg2 (the murine homolog of 
human ABCG2), or both [525, 526]. Interestingly, these studies suggested cooperativity 
between the two transporters in keeping substrates out of the brain. In the case of 
vemurafenib, for example, deletion of Abcb1a/b and Abcg2 resulted in a nearly 20-fold 
increase in the brain concentration of vemurafenib 24 hours after a 5 mg/kg dose 
compared to wild-type mice, while deletion of Abcb1a/b or Abcg2 alone resulted in 
virtually no increase [527]. Despite the growing evidence that these transporters remain 
a significant impediment to the treatment of brain metastases, no therapies have been 
approved to increase penetration of chemotherapy drugs into the brain. While mouse 
models have provided many insights into the roles of ABC transporters at the BBB, 
mice are expensive to maintain and are not amenable to high-throughput screening. 
Thus, the development of models that would lead to novel therapies to treat brain 
metastases would be of importance in the treatment of cancer. 
The zebrafish (Danio rerio, Dr) has emerged as a cheaper and more efficient 
model compared to rodents for studying the role of transporters at the BBB [528, 529]. 
Similar to humans, zebrafish have a functioning BBB comprised of endothelial cells 
that form claudin-5 and zona occludens-1 tight junction complexes that limit drug 
transcytosis [530, 531]. However, zebrafish possess no direct homolog of human 
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ABCB1; they instead have 2 genes that are closely related: zebrafish Abcb4 and 
Abcb5[532].  In the case of human ABCG2, there are 4 homologs: zebrafish Abcg2a, 
Abcg2b, Abcg2c and Abcg2d [533]. Our lab has previously characterized the different 
zebrafish homologs of human ABCB1 and ABCG2, and demonstrated that Abcb4 
[534] and Abcg2a have a substrate and inhibitor profile similar those of human 
proteins, respectively. More importantly, we discovered both Abcb4 and Abcg2a are 
present at the zebrafish BBB [534]. We have since been developing transgenic 
zebrafish models of the BBB where firefly luciferase or NanoLuc luciferase expression 
will be under the control of the glial fibrillary acidic protein promoter, leading to 
expression in the astrocytes behind the BBB. When the zebrafish BBB is disrupted 
either by inhibiting ABC transporters at the zebrafish BBB or otherwise increasing 
permeability, the luciferase substrates will cross the BBB and react with the luciferases, 
yielding a bioluminescent signal. Prior to a working zebrafish model, we must first 
determine if luciferase substrates are also ABC transporter substrates. The firefly 
luciferase substrate luciferin and the NanoLuc substrate coelenterazine are transported 
by ABCG2 or both ABCB1 [535] and Abcg2 [536], respectively. It is not known if 
luciferin, coelenterazine, or other coelenterazine analogs are substrates for the zebrafish 
transporters. In addition, we foresee the firefly luciferase zebrafish model having a 
much lower sensitivity compared to the NanoLuc luciferase model, as the NanoLuc 
luciferase reaction is known to be 150-fold brighter [537]. Here, we transfect human 
ABCB1 and ABCG2, and the zebrafish homologs Abcg4 and Abcg2a, as well as 
NanoLuc luciferase into HEK-293 cells to allow for complete characterization of the 
many NanoLuc substrates. The development of this zebrafish and cell-based model 
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could lead to the discovery of compounds that increase permeability of the BBB via 
novel mechanisms.  
8.2 Methods 
8.2.1 Cell Transfection 
HEK293 cells (ATCC, Manassas, VA) were transfected with empty pcDNA3.1 
vector or with vector containing full length abcb4 or abcg2a (Genscript, Piscataway, 
NJ) flanked by a FLAG tag. The MDR-19 and R5 cell lines were generated from 
HEK293 cells transfected with full-length human ABCB1 or ABCG2 and has been 
previously described [538]. Transfected cells were grown in MEM medium 
(Mediatech, Manassas, VA) and were maintained in 2 mg/ml G418 (Mediatech) and 
40 ?g/ml hygromycin B. Clones expressing similar levels of ABCB1, ABCG2, Abcb4 
or Abcg2a protein were selected based on FLAG and NanoLuc expression as detected 
by immunoblot.  
8.2.2 Immunoblotting 
Expression of ABC transporters and NanoLuc luciferase were examined using 
immunoblotting techniques. Transfected HEK293 cells were plated in 100-mm Petri 
dish (Falcon) at a density of 31,250 cells/cm2 for 24 hours. Whole cell lysates (20 ?g) 
were collected in RIPA buffer supplemented with 1% protease inhibitor and heated for 
20 minutes at 37?C. Protein lysates were separated on a 4-12% precast Bis-Tris gel 
(NuPAGE) and transferred onto a nitrocellulose membrane. After blocking with 
Odyssey blocking buffer (Li-COR) for 1 hour at room temperature, blots were probed 
with primary antibody against FLAG (Millipore-Sigma, Milwaukee), ABCB1 (C219, 
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Signet Laboratories, Dedham, MA, 1:250), ABCG2, or ?-actin overnight at 4 ?C. The 
blot was then incubated with IRDye 680RD goat anti-mouse secondary antibody (926-
68070, Li-COR) or IRDye 800CW goat anti-rabbit secondary antibody (926-32211, 
Li-COR) for 1 hour at room temperature based on the supplier?s advice. Visualization 
of protein bands was developed using the LiCor ODYSSEY CLx (Li-COR). ?-actin 
serviced as a loading control.  
8.2.3 Cytotoxicity Assays 
Transfected HEK293 cells were plated in 96 well plates (vendor) at a density of 
15,625 cells/cm2 and were allowed to adhere overnight. Cytotoxic drugs (doxorubicin 
for ABCB1& Abcb4; THZ1 for ABCG2 & Abcg2a) with or without ABC inhibitors (3 
?M elacridar for ABCB1 & Abcb4; 5?M Ko143 for ABCG2 & Abcg2a) were 
subsequently added in triplicate, and plates were incubated for 72 hours at 37 ?C. 
HEK293 cells not expressing a transporter (Empty vector cells, EV) was used as a 
control. Cell viability was measured via CellTiter-Glo (Promega, Madison, WI) assay. 
Luminescence was measured on a microplate reader (Tecan Infinite M200 Pro, Tecan 
Group) and GI50 (50% growth inhibitory) concentration values were calculated as the 
concentration at which 50% luminescence was observed compared to untreated cells.  
8.2.4 High-throughput Substrate Screening 
Transfected HEK293 cells were plated in 96 well plates at a density of 31,250 
cells/cm2. The next day, ABC inhibitors (3 ?M elacridar for ABCB1 & Abcb4; 5 ?M 
Ko143 for ABCG2 & Abcg2a) or complete media were added in quintuplicate and 
incubated with cells for 30 minutes at 37 ?C. Furimazine, coelenterazine, or 
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coelenterazine analogs, were subsequently added to the cells at a working solution of 
470nM for 1 minute at room temperature. Medium containing ABC transporters and 
NanoLuc substrates was immediately removed and replaced with fresh cell culture 
medium. After a 1-minute stabilization period, luminescence was measured on a 
microplate reader, every minute, for a total of 5 measurements. Brightness of the 
NanoLuc substrates was measured in EV cells in a similar manor but without the 
removal of the substrates.  
8.3 Results  
8.3.1 Establishment and Characterization of HEK293 Cells Stably Expressing an 
ABC Transporter and NanoLuc Luciferase 
To characterize NanoLuc substrate specificity with human ABCB1 and ABCG2, 
and zebrafish homologs, Abcb4 and Abcg2a, we transfected HEK293 cells with either 
empty vector plasmid, or vector encoding full-length ABCB1, abcb4, ABCG2, or 
abcg2a. Zebrafish transporter proteins both had a 5' FLAG tag. Clones were selected 
based on reactivity with an anti-FLAG, anti-ABCB1, anti-Abcg2, and anti-NanoLuc 
antibodies, as shown in Figure 8.1. As expected, the FLAG-tagged proteins ran slightly 
higher than the predicted molecular weight of zebrafish Abcb4 (141 kDa) and Abcg2a 
(72 kDa). Additionally, all transfected cells expressed low levels of human ABCB1, as 
HEK293 cells have a small baseline expression. 
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Figure 9.1 Immunoblot of HEK293 Transfected Cell Lines. HEK 293 cells overexpress 
human Abcg2 and ABCB1, zebrafish Abcg2a and Abcb4, and NanoLuc luciferase. 
 
8.3.2 Chemotherapy cytotoxicity Assessment within Double Transfected Cells 
To ensure the doubly transfected cell lines maintain their ABC transporter-
mediated efflux capabilities, cytotoxicity assays with known substrates of ABCB1, 
ABCG2, Abcb4, and Abcg2a were performed. Overexpression of human ABCB1 or 
zebrafish Abcb4 conferred resistance to doxorubicin (Figure 8.2b,c). Incubation with 
the elacridar inhibitor sensitized both cell lines to the chemotherapy, decreasing the 
GI50 by 47.9- and 23.0- folds, respectively. Similarly, the overexpression of human 
ABCG2 and zebrafish Abcg2a conferred resistance to THZ1, and the Ko143 inhibitor 
decreased the GI50 by 30.0- and 13.7- fold, respectively. As the zebrafish homolog 
proteins are not identical to the human transporters, it?s not unsurprising that the 
inhibitors were not as effective in reducing the GI50 (summarized in Table 1) for the 
zebrafish transfected cell lines. Alternatively, EV cells not expressing an ABC 
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transporter were sensitive to both chemotherapeutics (Figure 8.2a,d). EV cells 
incubated with doxorubicin and with or without the elacridar inhibitor, had a GI50 of 
approximately 0.0285 ?M. Similarly, EV cells incubated with THZ1, with or without 
the Ko143 inhibitor, had a GI50 of approximately 0.0255 ?M.  
Figure 9.2 Cytotoxicity Screening within Transfected HEK 293 cells. Double transfected 
cell lines maintain ABC transporter function. Transfected HEK293 cells were incubated with 
(a-c) doxorubicin with or without the elacridar inhibitor, or (d-f) THZ1 with or without the 
Ko143 inhibitor, for 3 days. N ? 6, where N equals the number of wells assessed over 3 trials. 
 
 
Table 9.1 GI50 Summary of Transfected HEK293 cells. GI50 values for (a) doxorubicin and 
(b) THZ1. 
 
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8.3.3 High Throughput Screening of NanoLuc Substrates 
The zebrafish larva brain at 5 days post fertilization is only ~1.5 mm in length 
and ~500 ?m in thickness [539]. Foreseeing sensitivity problems within the future in 
vivo assay, we first determine which NanoLuc substrates will produce the brightness 
luminescence signal. A panel of coelenterazine analogs were evaluated within EV cells. 
Furimazine, the substrate specifically developed to react with NanoLuc luciferase 
produce the brightness luminescence signal which was 35.9-fold larger than native 
coelenterazine (Figure 8.3). In fact, 4 other coelenterazine analogs produced a greater 
signal than native coelenterazine. The substrates that produced the smallest signal, 
coelenterazine-cp and coelenterazine-n, had a luminescence value that was 99.4% and 
97.3% smaller than furimazine.  
Figure 9.3 Bioluminescence Signal of Native Coelenterazine and its Analogs. HEK 29 cells 
that were transfected with NanoLuc luciferase but not an ABC transporter were incubated with 
a panel of luciferase substrates. N ? 6, where N equals the number of wells assessed over 3 
trials. 
 
Next, we determined if the NanoLuc substrates are also substrates for ABC 
transporters. Native coelenterazine and the 4 analogs that produced the largest 
bioluminescence signal (furimazine, coelenterazine-h, coelenterazine- hcp, 
coelenterazine-f) were screen in the panel of transfected cells. The bioluminescence 
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signal was measured every minute over a 5-minute period and the area under the curve 
was calculated (Figure 8.4). Native coelenterazine was a positive control as it was 
previously determined to be a substrate for both human ABCG2 and ABCB1. As seen 
in Figure 8.4a, native coelenterazine was a substrate for all 4 ABC transporters. The 
addition of an ABC inhibitor did not impact the bioluminescence signal produced by 
native coelenterazines within EV cells, but significantly increased the signal within all 
4 transfected lines. Furimazine, which produced the largest bioluminescence signal in 
EV cells, was determined to be a substrate for both human ABCG2 and zebrafish 
Abcg2a, but not ABCB1 nor Abcb4 (Figure 8.4b). Similarly, coelenterazine-h was 
only a substrate for ABCG2 and Abcg2a (Figure 8.4c). Alternatively, coelenterazine-
f was only a substrate for ABCG2 (Figure 8.4d). Lastly, coelenterazine-hcp was only 
not a substate for ABCB1 (Figure 8.4d). All NanoLuc substrates that screened were 
substrates for human Abcg2, while the majority were not substrates for human ABCB1. 
Similarly, the NanoLuc substrates were more likely to be a substate for zebrafish 
Abcg2a compared to Abcb4.  
 
 
 
 
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Figure 9.4 ABC Substrate Screening. A panel of transfected cells were incubated with (a) 
native coelenterazine and (b-e) coelenterazine analogs Substrates (c, f, h, f, or hcp) with and 
without ABC transporter inhibitors (+ i) were incubated with cells over expressing the 4 
different ABC transporters for 5 minutes. The area under the curve of the bioluminescence 
signal was calculated from 5 different time points. N ? 6, where N equals the number of wells 
assessed over 3 trials. 
  
While it?s important to determine which NanoLuc substrates are ABC 
transporter substrates, its equally important to determine the degree of which they are 
effluxed. Although a substate may produce a large bioluminescence, if it is a weak 
ABC substate, it can lower the specificity of the in vivo model. NanoLuc substrates 
that are not substrates for the ABC transporters will have a lower percent increase in 
bioluminescence near 0%, while strong substrates that are largely effluxed from the 
cell will have a percent increase in bioluminescence greater than 100% (Figure 8.5). 
EV cells incubated with all the NanoLuc substrates with or without an ABC inhibitor 
had a ~0% increase. The largest percent increase in bioluminescence occurred when 
coelenterazine-hcp was incubated with cells over expressing ABCG2 and the Ko143 
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inhibitor, at ~729.8?405.7% (Figure 8.5e). The largest percent increase for zebrafish 
Abcg2a and Abcb4 were coelenterazine-h at 97.3?45.7% (Figure 8.5c) and 
coelenterazine-hcp at 338.7?180.3%, respectively (Figure 8.5e).  
Figure 9.5 Bioluminescence Signal Percent Increase. Transfected HEK293 cells were 
incubated with (a) native coelenterazine or a (b-e) coelenterazine analog with and without an 
ABC transporter inhibitor (Ko143 for ABCG2 and Abcg2a; elacridar for ABCB1 and Abcb4). 
A single bioluminescence measurement at 5 minutes post incubation was used to calculate 
percent increase. N ? 6, where N equals the number of wells assessed over 3 trials. 
8.4 Discussion and Conclusions 
Treatment options for diseases of central nervous system remain limited [540]. 
Most therapeutics cannot effectively enter the brain on their own because of the intact 
BBB [541]. The endothelial cells of the BBB form tight junctions that restrict the 
passage of molecules [95]. Additionally, at the apical surface, the ABC transporters 
ABCB1 and ABCG2 work in concert to transport substrates back into the bloodstream 
and prevent access to the brain [542]. A number of chemotherapeutic agents are 
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substrates for one or both of the transporters.  This includes traditional chemotherapies 
such as paclitaxel, vinblastine, etoposide, colchicine and doxorubicin but also targeted 
therapies such as erlotinib, lapatinib and vemurafenib that have been approved to treat 
metastatic cancers of the lung, breast or melanoma, respectively [543-545].  Despite 
the clear need to develop treatment modalities to increase brain penetration of approved 
therapies, little progress has been made. 
Zebrafish offer a unique model to study transporter activity at the BBB. Our lab 
has recently discovered zebrafish Abcg2a and Abcb4 [534] transporters are present at 
the BBB and function similarly to their human homologs, ABCG2 and ABCB1, 
respectively. We are currently developing a transgenic zebrafish model that relies on 
NanoLuc luciferase luminescence to study the functions of these ABC transporters and 
BBB permeability. Once established, we see the potential for this model to be used for 
many applications that would impact public health by increasing our understanding of 
this critical physiological barrier and increasing brain penetration of cancer therapies. 
The transgenic fish could be used in high-throughput screens for inhibitors of ABCB1 
and ABCG2 at the BBB and would directly measure brain penetration of a small 
molecule caused by transporter inhibition. To our knowledge, this would be the first 
high-throughput assay to measure BBB penetration in a living organism. One of the 
major advantages of this system is that the transgenic fish can be used to identify not 
only direct inhibitors of transporter proteins at the BBB, but any compounds that might 
increase permeability of the BBB to small molecules.  For example, adenosine A2A 
receptor agonists have been shown to modulate transporter expression at the BBB 
without directly inhibiting the transporters [546]. The zebrafish model proposed here 
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would have utility in identifying similar compounds, unlike cell-based assays that can 
only measure direct inhibition. Additionally, this model could be used to assess the 
ability of novel methods?such as nanoparticles?to deliver chemotherapy to the brain. 
However, prior to a working zebrafish model, it is necessary to determine if NanoLuc 
luciferase substrates are also substrates for the zebrafish ABC transporters present at 
the BBB.  
Native coelenterazine and its analogs contain an imidazopyrazinone core, a 
nitrogen-bearing heterocycle, and three side groups [547]. Although there are only 
slight differences within the side groups corresponding to different amino acid side 
chains, the panel of coelenterazine analogs produced bioluminescence signals that were 
up to 155.7-fold different from each other. The zebrafish larva brain at 5dpf is 
extremely small, limiting the number of astrocytes that would express Nanoluc 
luciferase. Foreseeing sensitivity problems within the in vivo model due to a small 
bioluminescence signal, we first tested a panel of NanoLuc substrates within EV cells. 
As expected, cells incubated with furimazine, a substrate engineered to react with 
NanoLuc, produced the largest luminescence. Cells incubated with native 
coelenterazine produced a luminescence that was 35.9-fold smaller than furimazine. 
High-throughput screening experiments revealed all the NanoLuc substrates tested 
were also substrates for human ABCG2. Only native coelenterazine was a substrate for 
human ABCB1. More importantly, we identified that native coelenterazine, 
furimazine, coelenterazine-h, and coelenterazine-hcp were substrates for zebrafish 
Abcg2a, while coelenterazine-f was not a substrate. Similarly, native coelenterazine 
and coelenterazine-hcp were substrates for zebrafish abcb4. Using different Nanoluc 
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substrates (which are also ABC transporter substrates) within the in vivo zebrafish 
model could lead to differences in sensitivity and specificity when screening unknown 
ABC inhibitors. The largest percent increase in bioluminescent for zebrafish Abcg2a 
and Abcb4 were coelenterazine-h at and coelenterazine-hcp. Although furimazine 
produced the largest bioluminescent signal, other coelenterazine analogs that are 
stronger ABC transporter substrates may be more helpful in understanding BBB 
permeability.  
In Summary, the development of a transgenic zebrafish can increase our 
understanding of drug delivery to the brain. Identifying which luciferase substrates are 
effluxed by ABC transporters present at the zebrafish BBB is the first step to 
engineering this novel model. 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
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10. Chapter 9. Conclusions and Contributions to Science 
 
The invasive GBM cells that invade outwards into the healthy brain play a 
major role in disease recurrence. A vast amount of research has been dedicated to 
figuring out methods to eradicate these invasive cells. The fluorescence guided 
resection of the primary tumor has extended patient survival but does not adequately 
address the invasive nature of the disease. The intact BBB limits the ability of 
chemotherapy to accumulate in the invasive GBM cells and kill them. Numerous 
methods have been developed to open the BBB for the enhanced delivery of 
chemotherapy, but none are universally available in the clinic. Many of these BBB 
opening methods are lack specificity, have low efficacy, or can induce healthy tissue 
damage. PDT using the photosensitizer ALA-induced PpIX has shown promise in 
opening the BBB and treating GBM. However, the limited activation depth of PpIX in 
the brain stops the treatment from reaching the majority of the invasive cancer cells. In 
addition, large and non-optimized PDT doses within the brain can also result in healthy 
tissue damage. Here, we determined the activation depth of the clinically relevant VP 
photosensitizer in the rodent brain. We examined the mechanisms behind low dose 
PDT-mediated BBB opening in a transwell model. Furthermore, we engineered a novel 
pure-drug carrier-free nanoparticle of VP for improved cancer cell uptake and PDT 
treatment. We examined both PDT and PDP treatment within a subcutaneous GBM 
mouse model and intracranial rat model. Finally, we aided in the development of a high 
throughput zebrafish model that will allow us to study BBB integrity in mor detail.  
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9.1 Verteporfin can be Activated Up to 2 cm in the Rodent Brain 
 
FGR of GBM is routinely utilized in clinical centers around the world, whereas 
PDT treatment for GBM has yet to reach universal clinical implementation and remains 
an active area of preclinical and translational investigations. One of the major 
challenges to clinical use of PDT for brain cancer is the limited light penetration depth 
in brain tissues. To address this shortcoming, in Chapter 3, we developed Liposomal 
protoporphyrin IX and Liposomal verteporfin and evaluated their photodynamic effects 
as a function of depth in tissue and light fluence using rat brains. Although red light 
penetration depth (defined as the depth at which the incident optical energy drops to 
1/e, ~37%) is typically a few millimeters in most tissues, we show that the remaining 
optical energy could induce PDT effects up to 2cm into brain tissues. Photobleaching 
and singlet oxygen yield studies suggest that deep-tissue PDT (>1cm) is more effective 
when using Liposomal VP, compared to Liposomal PpIX. Together, these findings 
suggest that Liposomal VP based PDT is more likely to generate cytotoxic effects deep 
within the brain and offer the possibility of treating brain invading tumor cells 
centimeters away from the main, resectable tumor mass. Fluorescence imaging 
revealed ALA-induced PpIX remains the ideal choice for the FGS of GBM.  
9.2 PDP-induced Blood-brain Barrier Opening Enhances Drug Delivery 
  
The BBB remains a major obstacle for drug delivery to the central nervous 
system. In particular, the tight and adherens junctions that join the brain capillary 
endothelial cells limit the diffusion of various molecules from the bloodstream into the 
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brain. PDP is a non-cytotoxic modality that involves light activation of photosensitizers 
to photochemically modulate nearby molecules without killing the cells. In Chapter 4, 
we investigated the effects of sub-lethal photochemistry on junction phenotype (i.e., 
continuous, punctate, or perpendicular), as well as the BBB permeability in a transwell 
model of HBMECs. We showed that PDP decreases the mature continuous junction 
architecture by ~20%, increases the immature perpendicular junction architecture by 
~40%, and has minimal impact on cell morphology in HBMECs. Furthermore, 
transwell permeability assay revealed that PDP improves the HBMEC permeability to 
dextran or nanoliposomes by up to 30-fold for 6-9 days. These results suggest that PDP 
could safely reverse the mature brain endothelial junctions without killing the 
HBMECs. This study not only emphasizes the critical roles of PDP in the modulation 
junction phenotype, but also highlights the opportunity to further develop PDP-based 
combinations that opens the cerebrum endothelium for enhanced drug transporter 
across the BBB. 
 
9.3 Combinations of Photosensitizing Biomolecules may increase PDT Efficacy, but 
Limits Cellular Accumulation 
Many photosensitizers including VP are hydrophobic and require a drug 
delivery vehicle for patient administration. Photosensitizing biomolecules (PSBM) 
represent a new generation of light-absorbing compounds that could improve optical 
and physicochemical properties for biomedical applications. Despite numerous 
advances in lipid-, polymer-, and protein-based PSBMs, their effective use requires a 
fundamental understanding of how macromolecular structure influences the 
physicochemical and biological properties of the photosensitizer. In Chapter 5, we 
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prepared and characterized three well-defined PSBMs based on the photosensitizer, 
VP. The PSBMs include 16:0 lysophosphocholine-VP (16:0 Lyso-PC-VP), 
distearoylphosphoethanolamine-polyethylene-glycol-VP (DSPE-PEG-VP), and anti-
EGFR cetuximab-VP (Cetuximab-VP). In two glioma cell lines, DSPE-PEG-VP 
exhibited the highest singlet oxygen yield but was the least phototoxic due to low 
cellular uptake. 16:0 Lyso-PC-VP had the highest PDT efficacy but redirected VP?s 
subcellular localization from mitochondria to lysosomes. At 24 hours after incubation, 
proteolysed Cetuximab-VP was localized to mitochondria and effectively disrupted 
the mitochondrial membrane potential upon light activation. Our results reveal the 
cellular uptake, variable trafficking, and end effects of PSBMs, provide valuable 
insights into methods of PSBM evaluation, as well as strategies to select PSBMs based 
on subcellular targets and cytotoxic mechanisms. We ultimately demonstrated that 
PSBMs reduced cancer cell uptake and PDT killing compared to free-form VP, 
however, biologically informed combinations that target different organelles, 
concurrently, may lead to enhanced therapeutic effects. 
9.4 Pure-drug Carrier-free Nanoparticles of Verteporfin have Superior PDT Efficacy 
Compared to Liposomes 
 
A variety of photosensitizers have been investigated and marketed for PDT of 
cancer. Since the FDA approved the first and only nanoformulation of 
photosensitizer?Liposomal VP?for PDT of macular degeneration 20 years ago, 
significant advancements have occurred in our understanding of photosensitizer 
delivery. While liposomes significantly increase the ability of water-insoluble 
photosensitizers to achieve a clinically relevant concentration in aqueous media, 
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liposomes enclosed by phospholipid bilayers could also hinder the uptake and 
photochemical activity of photosensitizers in cells. In Chapter 7, we report a facile 
surfactant-free method to prepare amorphous nanodrug of VP (NanoVP) with 100% 
active pharmaceutical ingredients and nearly 1,500-fold higher photosensitizer-loading 
capacity compared to standard Liposomal VP. Monodispersed NanoVP is self-
quenched during storage in saline solution and can be de-quenched in cells, enabling 
PDT and fluorescence imaging of cancer cells. We demonstrate up to 10-fold 
enhancement in anti-tumor PDT efficacy of NanoVP compared to Liposomal VP in 
GBM cells and mouse models. Importantly, we also reported that low dose NanoVP-
PDT priming can be used for safe, localized, and controlled BBB opening, increasing 
Evans Blue accumulation in rat brains by 5.5-fold, compared to using FDA-approved 
ALA-induced PpIX photosensitizer. Collectively, the NanoVP represents a next-
generation versatile photosensitizer formulation and may facilitate treatment strategies 
for brain tumors and invasive central nervous system diseases that are protected by the 
intact BBB. 
9.5 A High Throughput Zebrafish Model aids in the Study of BBB Integrity 
The BBB represents a significant impediment to the treatment of brain cancers 
and cancers that have metastasized to the brain.  Brain diseases are so intractable due 
to, in part, the expression of ABC transporters at the BBB.  The ABC transporter family 
is comprised of 48 members that use ATP to transport substrates against a concentration 
gradient.  Two transporters that are found at very high levels at the BBB are P-gp (or 
ABCB1, encoded by the ABCB1 gene) and the ABCG2 (encoded by the ABCG2 gene).  
Counted among the substrates of P-gp and ABCG2 are a number of traditional 
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chemotherapeutic agents including paclitaxel, vinblastine, vincristine, doxorubicin and 
daunorubicin as well as a number of targeted therapies such as erlotinib, lapatinib and 
vemurafenib [110].  
Knockout mouse models have consistently demonstrated the major role ABC 
transporters play in keeping chemotherapeutic agents out of the brain. However, mouse 
models cannot be used to perform high-throughput studies. Recent reports have 
suggested that the Zebrafish (Danio rerio) BBB is a useful surrogate for the human 
BBB. The zebrafish BBB contains claudin-5 tight junction complexes and ABC 
transporter homologs that restrict many of the same chemotherapeutics from entering 
the brain. In Chapter 8, we develop a high throughput cell-based assay to study 
NanoLuc luciferase and ABC transporter substrates. Screening revealed that all 
coelenterazine analogs tested were substrates for human ABCG2 and the majority were 
substrates for zebrafish Abcg2a. Only 2 coelenterazine analogs were substrates for 
zebrafish Abcb4.  Foreseeing sensitivity problems with the zebrafish model, we 
determined that furimazine produced a 35.9-fold larger bioluminescent signal than 
native coelenterazine. Determining if luciferase substrates are also ABC transporter 
substrates is a critical first step in the development of a transgenic zebrafish model. We 
see the potential for this model to be used for many applications that would impact 
public health by increasing our understanding of this critical physiological barrier and 
increasing brain penetration of cancer therapies. 
9.6 Closing Remarks  
Overall, this dissertation advocates for the addition of NanoVP-mediated PDP 
into the clinical workflow for GBM. Evidence suggests that NanoVP-mediated PDP 
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can open the BBB for the enhanced delivery of drugs to the brain. Incorporating this 
BBB opening method can allow for a dose reduction of TMZ as more will be able to 
reach the invasive cells. In addition, other chemotherapies that normally can?t cross the 
BBB could then be used. NanoVP-mediated PDT can also be used for eradicating 
remaining GBM cells after surgery. Given that the feasibility of PDT has already been 
demonstrated in the clinic for the treatment of GBM and that fluorescence guided 
resection already relies on the necessary equipment, leveraging NanoVP for PDP and 
PDT treatments can help address the challenges associated with standard 
chemotherapy. Further investigations into multicycle treatments are warranted to fully 
exploit NanoVP as a PDT and PDP agent. 
9.7 Contributions to the Field 
9.7.1 Scientific Contributions 
The work in this dissertation makes the following contributions to the understanding 
of drug delivery and PDT: 
? Established that 690nm light activation of VP can occur at deeper depths, up to 
2 cm, in the rat brain compared to 635nm light activation of PpIX (Chapter 3); 
? Established that light activation of VP results in a ~1.7-2.5-fold higher 
production of 1O2 than PpIX in situ (Chapter 3); 
? Established that low dose VP-mediated PDT increases the permeability of 
100nm liposomes in a transwell model without killing the endothelial cells 
(Chapter 4); 
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? Established that low dose VP-mediated PDT increases the amount of immature 
perpendicular cell-cell junction and simultaneously decreases the amount of 
mature continuous junction (Chapter 4); 
? Established that covalent conjugation of VP to three different biomolecules 
decreases cellular uptake compared to free-form VP (Chapter 5); 
? Established that total VP uptake better predicates PDT efficacy than in situ 1O2 
production (Chapter 5); 
? Established that dual targeting mitochondria and lysosome during PDT 
treatment can result in enhanced cell killing (Chapter 5); 
? Established that pure-drug nanoparticles of VP (NanoVP) have superior PDT 
efficacy than Liposomal VP as a direct result of increased cell accumulation 
(Chapter 7); 
? Established that low dose NanoVP-mediated PDP can open the rat BBB 
without causing major healthy tissue damage (Chapter 7); 
? Established that the majority of coelenterazine analogs were substrates for 
human ABCG2 and zebrafish Abcg2a while only one was a substrate for 
human ABCB1 and two were substrates for zebrafish Abcb4 (Chapter 8); 
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Table 10.1. Summary of Contributions to the field. 
Chapter Considerations Current methods Gaps in the field Our current contribution
Comparing the activation depth of two 
Activation depth of Necrosis (histology staining after PDT Labor and time intensive. PS?s using an ex vivo model.
3
photosensitizers treatment), photon detection Only focus on light not PS. (Inglut C., Gaitan B., et al., Photochem. 
and Photobio. 2020)
BBB opening (osmotic solutions, Using low doses of PDT, known as priming, 
ultrasound, targeted nanoparticles), Low efficacy, invasive, to modulate tight junctions.
4 BBB modulation
Convection-enhanced, Interstitial healthy brain damage (Inglut C., Gray K. et al, IEEE J Sel Top 
delivery Quantum Electron. 2020)
Development of a pure drug nanoparticle 
Photosensitizer & understanding liposome design PDT for GBM relies on pro-drug, ALA. Long term GBM survival has 
parameters.
5-7 delivery & PDT Gold standard to deliver VP is via not been achieved with 
(Inglut C., Baglo Y., et al, J Clin Med. 2019)
applications liposomes either formulation. (Inglut C., et at., Nanomaterials. 2020)
(Inglut C., et al., Submitted)
Mice are expensive and total 
A zebrafish model to examine ABC 
Mouse model, cell culture model, organ variables are limited. Cell 
8 BBB modeling transporter substrates and inhibitors.
on a chip culture doesn?t accurately 
(Inglut C., et al., In preparation)
represent BBB.
 
9.7.2 Peer-Reviewed Journal Publications 
 
Published manuscripts: 
 
? Inglut C.T., Goodfriend (Gray) K.M., Vig S., Jung J.W., Stabile J., Zhang Y., 
Stroka K.M., Huang H.C. ?Photodynamic Priming Modulates Endothelial Cell?
Cell Junction Phenotype for Light-activated Remote Control of Drug Delivery? 
IEEE JSTQE. 2020.  
? Inglut C.T., Sorrin A.J., Kuruppu T., Vig S., Cicalo J., Ahmad H., Huang HC. 
?Immunological and Toxicological Considerations for the Design of 
Liposomes? Nanomaterials. 2020. 
? Gaitan B., Inglut C.T., Liu Y., Chen Y., Huang H.C. ?Depth Resolved Imaging 
of Photosensitizer in the Rodent Brain using Fluorescence Laminar Optical 
Tomography? J. Biomed. Opt. 2020. 
? Goodfriend (Gray) K.M., Jung J.W., Inglut C.T., Huang H.C., Stroka K.M. 
?Quantitatively Relating Brain Endothelial Cell-Cell Junction Phenotype to 
204 
 
 
Global and Local Barrier Properties Under Varied Culture Conditions via the 
Junctional Analyzer Program? Fluids and Barriers of the CNS. 2020. 
? Inglut C.T.+, Baglo Y. +, Liang B.J., Cheema Y., Stabile J., Woodworth G.F., 
Huang H.C. ?Systematic Evaluation of Light-Activatable Biohybrids for Anti-
Glioma Photodynamic Therapy? J. Clin. Med. 2019. (+, equal contribution)  
? Inglut C.T. +, Gaitan B. +, Najafali D., Lopez I.A., Connolly N.P., Orsila S., 
Perttil? R., Woodworth G.F., Chen Y., Huang H.C. ?Predictors and Limitations 
of the Penetration Depth of Photodynamic Effects in the Rodent Brain? 
Photochem Photobiol. 2019. (+, equal contribution) 
? Stabile J., Najafali D., Cheema Y., Inglut C.T., Liang B.J., Vaja S., Sorrin A.J., 
Huang H.C. ?Engineering gold nanoparticles for photothermal therapy, surgery, 
and Imaging. In Nanoparticles for Biomedical Application - Fundamental 
Concepts, Biological Interactions and Clinical Applications? Elsevier: Chapter 
12. Chung EJ and Rinaldi C, editors. 2019. 
Submitted manuscripts: 
 
? Inglut C.T., Quinlan J., Rahman I., Stable J., Gaitan B., Srivastava P., Chiou 
W., Karim B., Connolly N., Robey R., Woodworth G., Gottesman M., Huang 
H.C. ?Carrier-free, amorphous verteporfin nanodrug for enhanced 
photodynamic cancer therapy and brain drug delivery? Submitted to Advanced 
Materials. 2022.  
? Gaitan B., Inglut C.T., Kanniyappan U., Xu H., Li L., Huang H.C., Chen Y. 
?Development of an endoscopic auto-fluorescent sensing device to aid in the 
205 
 
 
detection of breast cancer and inform PDT treatment?. Submitted to IEEE 
Transactions on Biomedical Engineering. 2022. 
Manuscripts in preparation:  
? Inglut C.T., Quinlan J., Robey R., Thomas J., Huang H.C., Gottesman M. 
?Development of a High-Throughput Screening Strategy for Discovery of 
Human and Zebrafish ABC Transporter Substrates?. In Preparation. 2022 
9.7.3 National and International Conference Presentations 
 
(* denotes presenter) 
 
? ?Clinically Relevant Pure Porphyrin Nanoparticles for Photodynamic Therapy 
and Priming Applications?. Inglut C*, Quinlan J, Rahman I, Stable J, Gaitan B, 
Srivastava P., Chiou W, Karim B, Connolly N, Robey R, Woodworth G, 
Gottesman M, Huang HC. SPIE Photonics Europe, Strasbourg, France, April 
2022. 
? ?Photodynamic Modulation of Bidirectional Drug Transport Across the Blood-
Brain Barrier?. Inglut C, Liang B, Huang HC*. ASP 2021 Symposium - 20th 
Anniversary for Theresa Busch. June 2021. 
? ?Photodynamic Therapy for Brain Cancer: Tissue Penetration and Blood Brain 
Barrier Modulation?. Inglut C*, Gaitan B, Gray K, Najafali D, Stabile J, Lopez 
IA, Jung J, Stroka K, Chen Y, Huang HC. 2019 BMES. Cancer Drug Delivery 
(Cancer Technologies; Drug Delivery), Philadelphia, PA, October 2019. 
? ?Photosensitizing Biomolecules and Nanocrystals for Anti-Glioma 
Photodynamic Therapy?. Inglut C*, Baglo Y, Liang BJ, Stabile J, Cheema Y, 
206 
 
 
Najafali D, Woodworth GF, Huang HC17th International Photodynamic 
Association World Congress, Cambridge, MA, July 2019. 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
11. Chapter 10. Future Work and Outlook 
 
The work in this dissertation has motivated several additional projects and new 
areas of research, many of which will contribute to a more comprehensive 
207 
 
 
understanding of drug delivery, BBB opening and modeling, and PDT treatment. For 
example, Chapter 5 focused on the development and characterization of VP-
biomolecules. While different combinations of VP-biomolecules have the potential to 
increase PDT efficacy, it was determined that covalent conjunction of VP to 
biomolecules decreased cellular uptake. Two additional research projects lead by 
fellow lab mates, Dr. Barry Liang and Dr. Yan Baglo, examined ways to overcome the 
downsides of the new VP-biomolecules. Their research focused on enhancing delivery 
of antibody-VP conjugates to cancer cells via the ?carrier effect? [548] and escaping 
ABC transporter-mediated cellular efflux using lipid-VP conjugates [196]. Several of 
the future projects that were motivated by the results introduced in this dissertation are 
discussed below and preliminary results are presented.  
10.1 The Treatment of Intracranial Brain Tumors 
 
A logical extension of the work presented in this dissertation is the treatment of 
brain tumors with NanoVP-mediated interstitial PDT. For interstitial PDT one or more 
laser fibers can be inserted into the brain, through the primary tumor or near the 
invasive GBM cells. Light can be delivered through the end of the fiber or throughout 
a fiber with a cylindrical diffuser. Interstitial PDT could help overcome light 
penetration concerns as the light source is now at the center of the tumor instead of the 
resection cavity edge. Additionally, interstitial PDT could be more useful than surface 
illumination PDT if MRI imaging reveals the majority of the infiltrative cancer cells 
are localized to one area/path. In a small-scale clinical trial that examined interstitial 
PDT for recurrent GBM treatment, 4 out of 10 patients survived over 24, when the 
expected survival is only 6-8 months [549]. 
208 
 
 
Preliminary studies of the treatment of intracranial brain tumors have been 
performed using NanoVP-mediated interstitial PDT. Here, U87 cells were implanted 
Figure 11.1 NanoVP-mediated Interstitial PDP Treatment does not Result in Healthy 
Tissue Damage. U87 cells were implanted into the rat brain and were treated with interstitial 
PDP 10 days later. 24 hours post treatment, brains were harvested and cryosection. N =1-2 
mice. 
into the brains of two rats. Approximately 10 days later when the tumor is expected to 
fill most of the right brain hemisphere, treatment was performed. Rats were 
administering a low dose of NanoVP (0.25 mg/kg) and 30 minutes later PDT (40 J, 100 
mW) was performed using a single laser fiber that was inserted into the same burr hole 
used for tumor implantation. A single rat that had the laser fiber inserted into the brain 
but didn?t receive PDT treatment was used as a control. After treatment, the incisions 
were closed, and rats were allowed to recover for 24 hours. The rats were then 
euthanized, and the brains harvested. Cryosections of the brain were stained with H&E 
or Luxol faster blue (methods described in Section 7.2.15). Results revealed the two 
treated animals had a tumor diameter of 5.4 mm2 and 45.3 mm2 while the control animal 
had a tumor diameter of 95.8 mm2. More importantly, no signs of brain damage 
surrounding the tumor were present in the PDT treated animals.  
209 
 
 
Future work for this project includes combining PDP and PDT. First interstitial 
PDP can be performed to open the BBB and enhance the delivery of NanoVP to the 
tumor. After varying VP accumulation via fluorescence imaging, a second PDT does 
can be performed to kill the remaining cancer cells. Additionally, a larger dose of 
NanoVP can be administered to exploit VPs chemotherapeutic effects (discussed in 
Section 7.3.3 and 10.4) 
10.2 PDP Synergizes with Irinotecan and Minocycline 
 
Irinotecan is an effective salvage chemotherapy for GBM patients with a 
compromised BBB. Irinotecan functions as a topoisomerase I (Top1) inhibitor, 
trapping the Top1 DNA cleavage complex which causes double-stranded DNA breaks 
and cell death. However, the full potential of irinotecan remains to be hindered by the 
inability to cross the intact BBB, the overexpression of DNA damage repair enzymes 
(i.e., tyrosyl-DNA phosphodiesterase 1, Tdp1 [550]) and normal tissue toxicity. A two-
prong strategy to overcome these challenges includes: (1) The selective and reversible 
opening of the BBB with non-cytotoxic PDP; and (2) The repurposing of minocycline, 
a Tdp1 inhibitor and a neuroprotective agent. Minocycline, a broad-spectrum 
antibiotic, has been shown to inhibit the expression of Tdp1, enhancing the potency of 
irinotecan [198]. The central hypothesis is that the photo-activatable delivery of 
mechanism-based combinations in a spatiotemporally controlled manner will 
supersede the blind administration of cytotoxic cocktails. 
Previous studies have shown the ability of PDP to synergize with irinotecan for 
the treatment of pancreatic tumors [175]. In addition, studies have shown that 
irinotecan and minocycline synergize for the treatment of ovarian cancer [198]. Here, 
210 
 
 
we examine the effectiveness of the triple combination for the treatment of GBM in 
vitro and in vivo. First, the three free-form drugs were tested in cell culture. U87 cells 
were plated and allowed to adhere overnight. The next day 12 ?M irinotecan, 30 ?M 
minocycline, and/or 0.25 ?M VP were added to the cell. On day 3, PDT (0.25 J/cm2, 
10 mW/cm2) was performed and on day 5, cell viability was assessed via MTT assay. 
Results show all dual therapies deceased cell viability more than monotherapies and 
the triple therapy modestly decreases cell viability compared to dual therapies, although 
only significantly more than minocycline + PDT (Figure 10.2). This is unsurprising as 
the dual therapies have already shown to be synergistic in other cancer types. 
Differences in cell viability can be further increased by optimizing the treatment 
timeline, as pretreatments with minocycline might be necessary to decrease the 
expression of Tdp1.  
150
**
n.s. n.s.
100
*
n.s.
50
0
ol Ir R no T no T T let In i D i D D po M P
i
c + 
M  P  P tr
I I 
+  +
IR IR
o
in
m
Figure 11.2 Triple Combination Treatment within U87 Cancer Cells. U87 cells were 
incubated with irionotecan (IRI), minocycline (mino), and/or VP (PDT) for 72 hours. n = 4, *P 
< 0.05, **P < 0.01, n.s.: nonsignificant. 
 
Knowing the dual therapies can well manage tumor growth, we next test the 
triple combination in an aggressive subcutaneous mouse model. Here, GBM39 tumors 
211 
 
Viability (%)
 
that were grown and continually passaged in the mouse flank were dissociated and 
implanted into new mice. Treatment began thirteen days after implantation, when 
tumors reached approximately 120 mm3 (Figure 10.3), minocycline (40 mg/kg) was 
orally administered for 12 consecutive days (days 13-24). On days 17 and 24, 
Liposomal VP (0.25 mg/kg) and Liposomal irinotecan (20 mg/kg) were intravenously 
administered, and 2 hours later, PDT (50 J/cm2, 50 mW/cm2) was performed. Results 
from this study indicate that the addition of PDP to irinotecan and minocycline 
treatment modestly decreases tumor volume (Figure 10.4). Compared to the PDT 
treatment performed in Section 7.3.4 that resulted in noticeable necrosis, PDP left no 
signs of damage. Due to the aggressive nature of this patient derived cell line and 
beginning the treatment regimen when tumors were already above 100 mm3, it is not 
surprising the triple combination did not effectively manage tumor growth. Usually, 
PDT and chemotherapy treatments are administered when the tumor is 50-100 mm3. 
Here, the fist cycle of PDT and irinotecan was administered when the tumors were 
~200 mm3.  
Future work for this project, being conducted by Sumioa Pang and Dr.  Payal 
Srivastava, includes identifying cancer types with a high Tdp1 and Top1 activity ratio 
and engineering a nanoparticle that contains all three active pharmaceuticals: VP, 
irinotecan, and minocycline.  
212 
 
 
GBM39 Subcutaneous mouse model
Minocycline
Irinotecan
PDT
Day 0 Day 13 14 15 16 17 18 19 20 21 22 23 24
(tumors implanted) (~120 mm3)
Figure 11.3 In ViMviono cTycrlineea: 4t0m megn/ktg  (Torailm gaevalgien) e. On day 13 post tumor cell implantation, animals 
Irinotecan: 20 mg/kg (IV, liposome)
received minocyclBiPnDe:  0(.4250 m mg/kgg/  k(IVg, ,li poorsaoml)e,) irinotecan (20 mg/kg, IV), and VP-mediated PDP (0.25 
mg/kg, 50 J/cm2, 50 mW/cm2). 
DLI: 2 hours
PDT: 50 mW/cm2, 50 J/cm2  
20
Control
15 IRI
Mino
10 IRi + Mino
IRI + Mino + PDT
5
0
0 5 10 15 20 25
Days post treatment
Figure 11.4 GBM39 Normalized Tumor Volume. Minocycline treatment began on day 0. 
PDT and irinotecan were administered on days 4 and 11. n=4-6 animals per group. Performed 
with help from Sumiao Pang. 
10.3 A Second-generation Targeted Version of NanoVP 
Active targeting of nanoparticles toward a desired site has been a main research 
topic in nanomedicine for decades. In 2021, Japan approved the first 
photoimmunoconjugate, Alluminox, which combines an anti-PD1 antibody with a 
photosensitizer for head and neck cancer. As mentioned in Chapter 5, targeted 
nanoformulations offer a unique solution to differentiate drug delivery between cancer 
cells and normal cells. Targeted nanoparticles can deliver toxic treatments 
preferentially to diseased cells. We previously reported the development of an EGFR 
antibody conjugated with VP (Cetuximab-VP) for GBM treatment. However, studies 
213 
 
Tumor Volume (Normd.) 
 
revealed VP uptake and photokilling were severely hindered. This new project, 
overseen by Idrisa Rahman, entails engineering a targeted formulation of NanoVP. The 
central hypothesis is delivering a high payload of VP to cancer cells via active targeting 
will increase PDT efficacy while minimizing healthy tissue damage. As stated in 
Section 7.3.1, we have created a stable NanoVP suspension coated with neutral polymer 
poloxamer 407. One goal of this project is to conjugate an EGFR antibody to the PEG 
end caps of poloxamer 407. Additionally, efforts are underway in creating a NanoVP 
particle that is partially composed of VP-NHS (as described in Section 5.2.1) for ligand 
conjugation.  
10.4 Exploring NanoVP as a Chemotherapeutic 
As mentioned in Section 7.3.3, high doses of VP can act as a chemotherapy 
without light irradiation. Recent clinical and preclinical studies have reported high 
doses of VP (?M-mM) can induce ?dark? toxicity in cancer cells including GBM 
(NCT04590664) [506-508]. A 10-100-fold higher dose of VP, compared to the PDT 
dose, has been shown to downregulate Bcl-2, disrupt the YAP/TAZ-TEAD complex 
interaction, and induce cancer cell death without light activation [506, 507]. This 
dissertation revealed NanoVP can deliver high payloads of VP to cancer cells for 
effective cell killing. Alternatively, Liposomal VP stimulated cancer cell death. A 
future project includes designing a sequential treatment plan that includes NanoVP-
mediated   PDP followed by large doses of NanoVP without light. The goal of PDP is 
to modulate the BBB and inhibit ABC transporters for enhanced drug delivery to cancer 
cells. The second dose of NanoVP will act as a YAP/TAZ inhibitor. The central 
hypothesis of the project is NanoVP-mediated PDP will increase delivery of VP via 
214 
 
 
BBB permeabilization and ABC transporter inhibition, resulting in enhanced VP-
mediated YAP sequestration effects in GBM cells. Preliminary experiments performed 
by John Quinlan show that this sequential treatment on GBM cells alone in vitro does 
not yield a synergistic or additive effect on cell killing (Figure 10.5). In this 
experiment, U87 cells are incubated with 0.25 ?M nanoVP prior to PDT (0-7.5J/cm2, 
10 mW/cm2). After light irradiation, fresh medium supplemented with 8-14 ?M 
NanoVP is added to the cells. Cell viability is measured via MTT assay 2 days later. 
Results show cell death is either driven by YAP inhibition (PDT ? 0.1 J/cm2) or by 
PDT (0.5 J/cm2), but an enhancement effect in vitro is not seen. Work is ongoing to 
evaluate the combination of these modalities in vivo to leverage PDP-mediated 
vascular permeabilization and ABC transporter inhibition to enhance drug delivery, 
enhancing YAP/TAZ sequestration effects.  
100
VP for YAP 
VP for PDT hv Sequestration
- x 14 mM
50 0.25 mM x 14 mM
- x -
0.25 mM x -
0
0 0.1 0.5 1 2.5 5 7.5
Fluence (J/cm2)
 
Figure 11.5 NanoVP-mediated PDP and YAP/TAZ Inhibition for GBM Cell Killing. U87 
cells were incubated with 0.25 ?M NanoVP prior to PDT, followed by 14 ?M NanoVP for two 
days. N=3. Conducted by John Quinlan. 
 
10.5 PDP can Modulate ABC Transporter Expression 
 
215 
 
Viability (%, normalized)
 
Our lab has previously shown VP-mediated PDT can directly damage ABC 
transporters and decrease ATPase activity, thereby increasing the intracellular 
concertation of drugs [111, 175, 551]. Free-form VP is a substrate for human ABCG2 
and ABCB1. When VP is within the ABC transporter binding pocket and PDT is 
performed, 1O2 can covalently crosslink the protein [551]. After creating and validating 
our transgenic zebrafish model, mentioned in Chapter 8, one goal is to examine the 
effects of PDT on BBB integrity and ABC transporter function. 
First, to determine if VP is also a substrate for zebrafish Abcg2a and Abcb4, 
transfected HEK 293 cells were incubated with VP, with and without an ABC 
transporter inhibitor, and the mean fluorescence intensity of VP was measured on 
FACs. As expected, the mean fluorescence ratio of VP to VP + inhibitor within EV 
cells (i.e., cells without a transporter) was 1.09, suggesting VP was not effluxed out of 
the cell. Similarly, the mean fluorescence ratio within ABCG2 and ABCB1 cells was 
0.291 and 0.566, respectively, suggesting VP was effluxed by both proteins. Here, we 
show the mean fluorescence ratio of VP within Abcg2a and Abcb1 was 0.58 and 0.417, 
respectively, suggesting VP is a substrate for both zebrafish proteins.  
Next, we examined the PDT efficacy within the transfected HEK 293 cells in 
order to determine a subtherapeutic dose for ABC transporter modulation experiments 
(Figure 10.6). Transfected cells were incubated with 0.25 ?M VP for 24 hours prior to 
PDT (0-4 J/cm2, 10 mW/cm2). As expected, EV cells were the least resistant to PDT 
treatment, having an IC50 of 0.167 J/cm
2. Cells overexpressing ABCG2, Abcg2a, 
ABCB1, and Abcb4 had an IC50 of 0.41, 0.35, 0.39, and 0.23, respectively. The 
216 
 
 
subtherapeutic dose of 0.1 J/cm2, which resulted in viability > 90%, was selected for 
modulation experiments. 
Preliminary experiments suggest PDP can modulate ABC transporter 
expression in transfected HEK293 cells (Figure 10.7). Here, cells overexpressing 
human ABCG2 were incubated with 0.25 ?M VP for 24 hours prior to PDP (0.1 J/cm2, 
10 mW/cm2). Cell lysates were collected, and protein expression was examined. 
Compared to the no treatment control, PDP modestly decreased the expression of 
ABCG2 to 0.95, 0.81, and 0.80 at 1-, 6-, and 24-hours post treatment, respectively. 
Although PDP treatment can decrease transporter expression by > 50% within cancer 
cells, these HEK293 cells are non-cancerous and were transfected.  
Future work for the project, being carried out by John Quinlan, includes using 
NanoVP-mediated PDP to modulate the ABC transporters at the zebrafish BBB and 
using coelenterazine analogs as permeability tracers to examine treatment outcomes.  
 
 
 150
 EV
 ABCG2
 100 Abcg2a
 ABCB1
 50 Abcb4
 
 
 0
 0.1 1 10
 Fluence (J/cm2)
Figure 11.6 PDT Dose Viability Curves. PDT was performed on HEK 293 cells transfected 
with NanoLuc luciferase and ABCG2, Abcg2a, ABCB1, and Abcb4. EV cells without a 
transporter were used as a control. N ? 6, where N equals the number of wells assessed over 3 
trials. 
217 
 
Viability (%)
 
 
 
 
 
 
 
Figure 11.7 Immunoblots of ABCG2 within Transfected HEK293 cells. VP-mediated PDP 
modestly decreases ABCG2 expression with transfected HEK293 cells. N=2 gels. 
10.6 A Zebrafish Model to Study Blood-brain Barrier Integrity 
As mentioned in Chapter 8, our goal is to develop a transgenic zebrafish model 
to study BBB integrity and ABC transporter activity. This ongoing project is currently 
being conducted by Dr. Joanna Thomas. The transgenic zebrafish model consists of 
NanoLuc luciferase expression under the control of the glial fibrillary acidic protein 
promoter, leading to expression in the astrocytes behind the BBB. When the zebrafish 
BBB is disrupted either by inhibiting ABC transporters at the zebrafish BBB or 
otherwise increasing permeability, the luciferase substrates will cross the BBB and 
react with the luciferases, yielding a bioluminescent signal. We have already shown 
that coelenterazine-h, a NanoLuc substrate, is a substrate for zebrafish Abcg2a within 
transfected HEK293 cells. Similarly, native coelenterazine is a substrate for Abcg2a as 
well as Abcb4 in HEK293 cells. Here, we show coelenterazine-h is transported by 
Abcg2a at the zebrafish BBB and produces a bright luminescence signal within the 
brain (Figure 10.8). Zebrafish at 7 days post feralization were preincubated with 10 
?M inhibitor for 2 hours prior to incubation with 3 ?M native coelenterazine or 
218 
 
 
coelenterazine-h. Luminescence was recorded every 15 minutes for 90 minutes. 
Without the addition of inhibitors, native coelenterazine and coelenterazine-h do not 
reach the NanoLuc containing astrocytes and produced a minimal luminescence signal. 
As expected, when Abcb4 is inhibited by valspodar and Abcg2a is inhibited by Ko-
143, native coelenterazine can interact with NanoLuc luciferase and produce 
luminescence, suggesting it is a substrate for both proteins. Similarly, when Abcg2a is 
inhibited by Ko-143, coelenterazine-h can cross the BBB and produce a significantly 
brighter luminescence, matching our in vitro data. Next steps for this project include 
examining wider concentration titrations of inhibitors to check dose response 
relationship and sensitivity of the assay. Our long-term goal is to test novel ABC 
transporter inhibitors and examine other ways to open the BBB within zebrafish. 
 
 
 
 
 
 
 
Figure 11.8 Native coelenterazine and coelenterazine-h are transported by Abcb4 or 
Abcg2a at the zebrafish BBB. Zebrafish (7 days post fertilization) were incubated with (a) 
native coelenterazine or (b) coelenterazine-h, with and without ABC inhibitors for 60 minutes. 
Native coelenterazine is a substrate for zebrafish Abcb4 or Abcg2a, and when the transporters 
are inhibited, it can cross the BBB and produce a luminescence signal. Coelenterazine-h is only 
a substrate for Abcg2a, and when it is inhibited by Ko-143, it can cross the BBB and react with 
NanoLuc luciferase. N ? 5, where N equals the number of fish. Conducted by Joanna Thomas. 
10.7 Outlook 
It is seemingly clear the GBM cells 1-2 cm away from the primary tumor edge 
underlie the near 100% disease recurrence. The intact BBB plays a significant role in 
219 
 
 
hindering current GBM treatments. Combining the proposed treatment techniques, 
NanoVP-mediated PDP, NanoVP-mediated PDT, and/or NanoVP-mediated YAP/TAZ 
inhibition, with the current treatment options hold significant promise in effectively 
treating GBM. Using the established ex vivo rat brain model and the proposed zebrafish 
model offer ways to better understand PDT treatment outcomes in the brain and the 
mechanisms hindering drug delivery to the brain. Furthermore, the expanded studies 
on an active targeting NanoVP formulation, planning sequential treatment strategies 
around NanoVP, and inhibiting ABC drug efflux transporters are the first steps in 
expanding NanoVP use for other disease types. In addition, the work in this dissertation 
has motivated the development of a triple combination nanoparticle/therapy to treat 
ovarian cancer. The knowledge presented in this dissertation has the potential to 
improve treatments for a wide range of cancers. 
220 
 
 
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