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Cellular uptake mechanism of elastin-like polypeptide fusion proteins

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Cellular uptake mechanism of elastin-like polypeptide fusion proteins

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CELLULAR UPTAKE MECHANISM OF ELASTIN-LIKE
POLYPEPTIDE FUSION PROTEINS

By

Xiaoli Pan

A Thesis Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
MASTER OF SCIENCE
(MOLECULAR PHARMACOLOGY AND TOXICOLOGY)

August 2018

2
ACKNOWLEDGEMENTS

I am really grateful to my mentor Dr. J. Andrew MacKay for his continuous
guidance and support throughout my whole master study. I would also like to
thank my thesis committee members, Dr. Curtis T. Okamoto and Dr. Roger F.
Duncan regarding their precious time and suggestions for my thesis. I am very
thankful to Santosh Peddi who trained me at the beginning, led me into the world
of science and provided many useful opinions throughout this project. Many
thanks to all other colleagues in MacKay lab for their wonderful help.

3
TABLE OF CONTENTS

ACKNOWLEDGEMENTS .................................................................................... 2
LIST OF FIGURES ................................................................................................ 5
ABSTRACT .............................................................................................................. 6
CHAPTER 1: INTRODUCTION ........................................................................... 7
CHAPTER 2: BACKGROUND ........................................................................... 10
2.1 Nanotechnology, nanomedicine and nanoparticle ...............................................................10
2.2 Elastin-like polypeptides ......................................................................................................11
2.3 Rapamycin, FKBP and mTOR signaling ...........................................................................15
2.4 Cellular uptake mechanism ................................................................................................18
CHAPTER 3: MATERIALS AND METHODS .................................................. 20
3.1 FKBP-ELP gene cloning ......................................................................................................20
3.2 FKBP-ELP expression and purification .............................................................................20
3.3 FKBP-ELP physicochemical characterization ...................................................................22
3.4 Fluorophore labeling ..........................................................................................................22
3.5 Rapamycin encapsulation and HPLC validation ...............................................................23
3.6 Dynamic light scattering ....................................................................................................24
3.7 Cell culture .........................................................................................................................25
3.8 Cold competition-binding assay and live cell imaging ......................................................25
3.9 Concentration-dependent cellular uptake ..........................................................................26
3.10 Rho-FAF, Fluorescein-dextran and Rhodamine B-dextran cellular uptake ....................26
3.11 Time-dependent cellular uptake and loss .........................................................................27
4
3.12 Immunofluorescence for mTOR targeting .......................................................................28
CHAPTER 4: RESULTS ...................................................................................... 29
4.1 Physicochemical characterization of FKBP-A192-FKBP .................................................29
4.2 Fluorophore labeling and rapamycin encapsulation ..........................................................30
4.3 Cellular uptake of FAF is receptor independent ................................................................32
4.4 Cellular uptake of FAF is through fluid-phase endocytosis and FAF goes to lysosomes as
the intracellular destination ..............................................................................................34
4.5 Cellular uptake and loss of Rho-FAF and Rho-FAF/Rapa are time dependent. .................38
4.6 Rho-FAF/Rapa targeting mTOR in MDA-MB-468 cells ..................................................42
4.7 Cellular internalization mechanism of FAF or FAF/Rapa .................................................44
CHAPTER 5: DISCUSSION ................................................................................ 45
CHAPTER 6: CONCLUSION ............................................................................. 48
REFERENCES ....................................................................................................... 49

5
LIST OF FIGURES
Fig. 1. Reversible phase separate behavior of ELPs ....................................................................12
Fig. 2. Schematic illustration of the purification technique for ELPs, termed Inverse Transition
Cycling (ITC) ...............................................................................................................................14
Fig. 3. The mechanism of action of rapamycin and the localization of mTORC1 ........................16
Fig. 4. Main pathways of endocytosis .........................................................................................19
Fig. 5. Physicochemical characterization of Rho-FAF ................................................................30
Fig. 6. Rhodamine labeling and rapamycin encapsulation cause no stability issues .....................31
Fig. 7. Ten-fold excess unlabeled FAF cannot compete off the signal of Rho-FAF suggesting no
receptor is involved upon binding ................................................................................................33
Fig. 8. Rho-FAF colocalizes with Fluorescein-dextran upon cellular uptake and the
internalization of Rho-FAF is not saturable ...................................................................................35
Fig. 9. Rho-FAF and Rho B-dextran show the same colocalization pattern with lysosomes ........37
Fig. 10. Time-dependent internalization profile of Rho-FAF with or without rapamycin ............39
Fig. 11. Cellular loss of Rho-FAF and Rho-FAF/Rapa .................................................................41
Fig. 12. Rho-FAF/Rapa targeting mTOR and mTOR localization in MDA-MB-468 cells ..........43
Fig. 13. Schematic of FAF/Rapa on cellular uptake pathways in MDA-MB-468 cells ................44

6
ABSTRACT
Rapamycin (Rapa) is a highly potent drug for preventing allograft rejection and cancer treatment
but has some drawbacks, including low solubility, poor bioavailability and cytotoxicity, which
limit its clinical potential. To improve Rapa delivery, our lab has developed a number of
nanocarriers by genetically fusing its cognate protein receptor, FKBP12, to a recombinant
protein-polymer, elastin-like polypeptide (ELP). ELPs have the amino acid sequence of
(VPGXG)
n
where X represents the guest residue and n specifies the number of repeats. One of
the FKBP-ELPs constructs, FKBP-A192-FKBP (FAF, X=Ala, n=192), outperformed other
carriers showing much higher tumor accumulation in an animal study. However, the cellular
uptake mechanism of this construct has not been elucidated and thus, leads to this study which
aims to investigate the internalization mechanism of FAF as well as its intracellular fates in
MDA-MB-468 cells. We found that no cell surface specific receptor was involved in FAF
binding. FAF colocalized with fluid phase endocytosis marker, dextran and the cellular uptake
had no saturation within 1-100 µM range, indicating that FAF was internalized through fluid
phase endocytosis. FAF targeted lysosomes as its intracellular destination and the cellular loss
half-life for FAF and FAF/Rapa were 17.7 hours and 21.3 hours respectively. mTOR was
colocalized with lysosomes in MDA-MB-468 cells; furthermore, FAF/Rapa also colocalized
with these structures after 1 hour. The results of this study provide new information about the
cellular uptake mechanism of ELPs and lay a basis for understanding how other ELPs fusion
carriers behave.
7
CHAPTER 1
INTRODUCTION
Recent developments of nanotechnology in medicine had a tremendous impact on drug delivery,
largely improving the performance of many existing drugs and facilitating the use of many new
therapies especially in the cancer field(LaVan et al., 2003). Among many applications,
nanoparticles have been studied very aggressively. Although there are many different kinds of
nanoparticles, people are particularly interested in biodegradable polymeric nanocarriers as they
may outperform others(Kumari et al., 2010; Soppimath et al., 2001).
Elastin-like polypeptides (ELPs) are one such kind of biosynthetic polymer that can fabricate
nanoparticles. They are a class of genetically engineered recombinant protein-polymer derived
from human tropoelastin, consisting of a pentapeptide repeats of (VPGXG)
n
. One of the most
interesting features of ELPs is that they are thermally responsive. At low temperature, they are
soluble in solution but can coacervate into an insoluble and ordered state upon being
heated(Meyer and Chilkoti, 1999). Besides the thermo-sensitive property, ELPs have many other
benefits such as biocompatible and biodegradable, precisely controllable and modifiable at the
genetic level, and can be easily and efficiently biosynthesized as monodisperse without involving
chemical polymerization or bioconjugation(Chilkoti et al., 2006). All these advantages render
ELPs as a potential tool in drug delivery and tissue engineering. ELPs were used to conjugate
with doxorubicin that can self-assemble into a sub- 100 nm nanoparticles, improving the
pharmacokinetics and lowering the toxicity of free drug(MacKay et al., 2009). Thermally
8
responsive ELPs can be designed to facilitate the application of local hyperthermia treatment to
the site of the tumor(Meyer et al., 2001). ELPs were fused to glucagon-like peptide-1 (GLP-1)
and designed to transit below the body temperature which can form a subcutaneously injectable
depot and lead to longer circulation time and a controlled release profile of GLP-1(Luginbuhl et
al., 2017).
Previously, our lab has successfully utilized ELPs to improve the performance of a small
molecule drug called rapamycin, also known as sirolimus. Rapamycin was developed as an
immunosuppressant in organ transplantation and more recently, its anticancer activity was
greatly being observed in many preclinical and clinical studies. Although it is an extremely
potent drug, it has a number of drawbacks like low solubility (~2.6 µg/mL)(Simamora et al.,
2001), poor bioavailability (14%-16%) and cytotoxicity (pulmonary- and nephron-toxicity).
With the goal to improve the solubility and toxicity profile of rapamycin, our lab genetically
fused ELPs with its native protein receptor FKBP (F) to facilitate the rapamycin delivery,
resulting in three different FKBP-ELPs constructs, FA (X=Ala, n=192), FAF (X=Ala, n=192)
and FSI [(X=Ser, n=48) + (X=Ile, n=48)](Dhandhukia et al., 2017; Shi et al., 2013). A series of
in vitro and in vivo studies were conducted to evaluate these three constructs, showing that all
three constructs retained rapamycin, suppressed tumor growth, affected the level of downstream
signaling, and increased tumor accumulation. In terms of first four aspects, these three constructs
performed similarly when given by intravenous administration. However, when given by
9
subcutaneous injection, FA and FAF accumulated to a higher level than FSI(Dhandhukia et al.,
2017).
Since we had very little cellular information about how ELPs directly interact with tumor cells,
thus we proposed to investigate the cellular uptake mechanism of FKBP-ELPs constructs, and
FAF was chosen to study. Understanding the fate of the drug carrier within the cell is very
important for drug delivery because it is necessary to explain how drug carrier releases the
encapsulated drug and how the drug binds its target in the cytosol. Thus, in this project, I
explored how FAF/Rapa goes into the tumor cells and what happens to render tumor suppression.
In all the studies, we chose to use MDA-MB-468 breast cancer cells because that was the cell
line used by our previous lab members to build the xenograft breast cancer mouse model to
evaluate the FKBP-ELPs constructs. FAF fusion protein was expressed and purified followed by
rhodamine labeling and rapamycin encapsulation. A cold competition-binding assay was
conducted to determine there was no specific cell surface receptor involved in FAF binding.
Dextran was used as a fluid phase endocytosis marker for comparison, and a
concentration-dependent cell uptake assay was carried out showing that FAF should be
internalized through pinocytosis. Lysosomes were the intracellular destination for FAF, and the
kinetic uptake and loss of FAF and FAF/Rapa in cells were monitored. Finally, mTORC1 was
confirmed to associate with lysosomes in MDA-MB-468 cells when activated under the
nutrient-rich condition, and FAF/Rapa was able to bind to it using immunofluorescence (IF).

10
CHAPTER 2
BACKGROUND
2.1 Nanotechnology, nanomedicine and nanoparticles
Nanotechnology is a rapidly developing and highly multidisciplinary domain, which promotes
the emergence of a massive and diverse array of excellent products derived from different fields
like engineering, chemistry, physics and biology(Mnyusiwalla et al., 2003). Its utilization to
medicine, “nanomedicine”, is currently under vigorous development for application in drugs,
drug delivery, in vivo imaging and diagnostics, biomaterials, active implant and etc.(Wagner et
al., 2006). The definition of nanotechnology has been proposed as the manipulation of matter
that is itself or has essential components in 1-100 nm range in at least one dimension(Porter et al.,
2008). However, more and more scientists emphasize less strict limitations on the exact
dimensions and describe the ‘right’ size(Whitesides, 2003) in bionanotechnology in a practical
way, including ‘micro-size’ (100 nm-1000 nm) systems with remarkable properties with respect
to addressable unmet medical needs (Ferrari, 2005).
Nanoparticles, primarily through the approval of various liposomal drug formulations, are among
the most successful applications in drug delivery. They have been extensively explored to
improve the delivery of small molecules, peptides, proteins or nucleic acids by employing their
unusual properties like the small size and the large surface to mass ratio (LaVan et al., 2003).
There are many potential advantages of using nanoparticles for drug delivery like increasing the
solubility of molecules, protecting molecules from metabolic degradation, providing active
11
targeting potential, enhancing drug uptake while reducing free drug toxicity, and achieving
controlled or sustained drug release(Singh and Lillard, 2009). Although all kinds of materials can
be used to fabricate nanoparticles (metal, silica, chemical synthetic or biosynthetic polymer,
peptides, lipids, etc.), more interests were put on those biodegradable materials especially
biosynthetic polymers because they can achieve effective drug release at the target sites without
introducing accumulation/toxicity from the drug carrier themselves (De Jong and Borm, 2008).
2.2 Elastin-like polypeptides
Elastin is a highly elastic protein in connective tissues and a key component of the extracellular
matrix (ECM) of lung, skin, blood vessels, ligaments, cartilage, tendon and etc. It imparts
vertebrate tissues elasticity and resilience to recover to their original shape and size after
contracting and stretching(Mithieux and Weiss, 2005). Elastin is the mature insoluble complex
that made by the lysine-mediated crosslinking of its soluble precursor, tropoelastin. Tropoelastin
is rich in hydrophobic domains dominated by alanine, valine, leucine, isoleucine and glycine.
The interaction between the hydrophobic domains can result in a phase separate behavior called,
coacervation(Sandberg et al., 1981). This process is reversible and thermodynamically controlled
as revealed by the previous study on partial hydrolysis product, α-elastin(Urry et al., 1969).
Elastin-like polypeptides(ELPs) are genetically-encoded recombinant protein polymer inspired
by the elastomeric domain of human tropoelastin. They are constitutive of a pentapeptide repeat
(VPGXG)
n
where X represents any guest amino acid and n specifies the number of pentapeptide
repeats(MacEwan and Chilkoti, 2014). ELPs follow the interesting feature of elastin that they
12
can undergo an inverse phase transition (Fig. 1a). They are soluble as random coils below a
transition temperature(T
t
), but coacervate into more ordered aggregate structure when heated
above the T
t
. This phase behavior happens in a short time and is typically reversible, meaning
that the coacervated ELPs can be redissolved when the solution is cooled down below

Fig. 1. Reversible phase separate behavior of ELPs. a, ELPs can change from a soluble state in
aqueous solutions to an insoluble coacervate state when being heated above the transition temperature.
b, The linear model fit shows that the transition temperature of ELPs is concentration dependent.
Three different ELPs with varied molecular weight and hydrophobicity exhibit distinct transition
temperature profiles.
Reproduced from Despanie et al., 2016 with permission of Elsevier.

13
the T
t
(Meyer and Chilkoti, 2004). The transition temperature can be tuned by many aspects (Fig.
1b) other than temperature, including internal factors like the hydrophobicity and molecular
weight of ELPs or external factors like ELPs concentration, salt concentration, solution pH and
etc. In brief, ELPs with higher molecular weight and more hydrophobicity guest residues display
lower transition temperature. Higher ELPs concentration and salt concentration also reduce
transition temperature (Despanie et al., 2016).
One major advantage of ELPs is that they can be biosynthesized in expression systems like
E.coli without any chemical synthesis and can be purified by exploiting their unique phase
transition behavior without complex chromatography. That largely reduces the cost of
manufacturing and makes laboratory scale-up production easier. The method used to purify ELPs
is called Inverse Transition Cycling (ITC) (Fig. 2). The purification process starts with cells lysis
using sonication and cellular debris removal followed by centrifugation. The supernatant is either
heated up above the T
t
or salts like NaCl are added to depress the T
t
below the solution
temperature to trigger the phase transition of the ELPs in the solution. The coacervate ELPs are
separated from the solution by centrifugation at the same temperature and the supernatant
containing soluble contaminants is discarded. This step is called a “hot spin”. In the next step,
termed the “cold spin”, the pellet containing ELPs from hot spin is resuspended in cold buffer
like PBS and then centrifuged at 4 °C to remove any insoluble contaminants while the
supernatant containing ELPs is decanted and retained. One hot spin and one cold spin together
14
are called one cycle of ITC. ITC usually needs to repeat multiple times to achieve the desired
ELPs with high purity(Hassouneh et al., 2010; Meyer and Chilkoti, 1999).
To expand the therapeutic usage of ELPs, some soluble proteins or peptides were genetically
fused with ELPs to improve their pharmacokinetics, biodistribution and therapeutic efficacy.
Interestingly, these recombinant fusion proteins can retain the thermal responsiveness property of
ELPs and thus, the purification can also be conducted following the ITC(Meyer and Chilkoti,
1999).

Fig. 2. Schematic illustration of the purification technique for ELPs, termed Inverse Transition
Cycling (ITC). ELPs or ELP fusion proteins can be separated and purified simply by using their phase
transition behavior which makes the process cost- and time- efficient and easy for scale-up.
Reproduced from Hassouneh et al., 2010 with permission of John Wiley and Sons.

15
2.3 Rapamycin, FKBP and mTOR signaling
Sirolimus (rapamycin, RAPAMUNE®) is a cyclic and hydrophobic macrolide antibiotic
produced by the bacterium Streptomyces hygroscopicus and was first isolated in 1972 from the
soil samples on Easter Island(VéZina et al., 1975). In the beginning, it was indicated as an
antifungal agent but its use was later shifted to an immunosuppressant in organ transplantation
due to the discovery of its immunosuppressive and anti-proliferative properties(MacDonald and
RAPAMUNE Global Study Group, 2001). In 2015, FDA approved another indication for
rapamycin, lymphangioleiomyomatosis (LAM), which is a very rare and progressive lung
disease. (McCormack et al., 2011). Interestingly, rapamycin was also found to extend lifespan
(Harrison et al., 2009), as well as anti-cancer activity including breast(Yu et al., 2001),
colon(Seeliger et al., 2004), prostate(Majumder et al., 2004) and kidney cancer(Luan et al., 2003).
Although the anti-tumor efficacy of rapamycin was extensively studied, its development into
clinical use was far slower due to the formulation and stability concerns in generating a
parenteral formulation(Bjornsti and Houghton, 2004). And combined with the drawbacks
mentioned before, more efforts needed to be done to successfully advance rapamycin as
anti-cancer drug to the market.
As for the mechanism of action of rapamycin, it first binds to its cognate receptor FK506 binding
protein-12 (FKBP-12) which belongs to the immunophilin family with a K
d
of 0.2 nM(Bierer et al.,
1990). The FKBP-rapamycin complex then associates with the FKBP-rapamycin binding domain
(FRB) of the mammalian target of rapamycin (mTOR), restricting the access or masking the
16
docking sites of mTOR substrates (Fig. 3a)(Benjamin et al., 2011; Hausch et al., 2013). There are
two structurally and functionally different mTOR complexes, mTORC1 and mTORC2. It is
thought that FKBP-rapamycin usually only inhibits mTORC1, but there is also research showing

Fig. 3. The mechanism of action of rapamycin and the localization of mTORC1. a, Rapamycin
firsts binds to its endogenous receptor FKBP12 and the binary complex then associates with
mTORC1. b, mTORC1 has two main localizations in cells. It usually associates with lysosomes when
it is activated under nutrient-rich condition, whereas distributed in the cytosol under starvation
inhibition state. c, Rapamycin-FKBP complex can inhibit mTORC1 signaling pathway which results
in its anti-proliferative property.
a and c are reproduced from Benjamin et al., 2011 with permission of Springer Nature.
b is reproduced from Betz and Hall, 2013 with permission of Rockefeller University Press.

17
that prolonged exposure of rapamycin can also have inhibition effect on mTORC2(Sarbassov et al.,
2006). The inhibition of mTORC1 pathway (Benjamin et al., 2011) causes the dephosphorylation
and inactivation of p70S6 kinase (Fig. 3c), resulting in the inhibition of ribosomal components
production which is essential for protein synthesis and cell-cycle progression. The process
ultimately blocks IL-2 stimulation of lymphocyte division which is the basis for rapamycin to
prevent allograft rejection(Guba et al., 2002; Kahan, 2000). In addition, increasing evidence
shows that mTOR is a central regulator of both cell growth and proliferation, as it can sense
nutritional status and mitogens in cells and act as a gatekeeper for progression from G1 to S
phase(Harris and Lawrence, 2003). Since in many cancers, mTOR pathway is highly upregulated
so it makes the strategy to inhibit mTOR in order to stop the dysregulation of G1 transit a very
potent target for cancer treatment(Bjornsti and Houghton, 2004).
The localization of mTORC1 in cells has also been studied (Fig. 3b). Interestingly, people found
that when being activated by amino acids or growth factors, mTORC1 is usually associated with
lysosomes but remains cytoplasmic and dispersed under amino acid starvation condition(Sancak
et al., 2010). Further exploration showed that mTORC1 is activated by GTP-bound Rheb on the
surface of the lysosomes. To achieve that, mTORC1 needs first translocate to the lysosomes
surface which is stimulated by nutrients and Rags. Rheb also needs to be activated, which
transforms from GDP- to GTP-bound form in response to growth factors(Betz and Hall, 2013).
The inhibition of mTORC1 by rapamycin does not affect mTORC1 localization to the lysosomes,
and also kinase-inactive mTORC1 still localizes with lysosomes, indicating that the activity of
18
mTORC1 does not affect its localization but may be related to its activation mechanism(Sancak
et al., 2008; Tabatabaian et al., 2010).
2.4 Cellular uptake mechanism
Cellular internalization of extracellular materials is a fundamental property of living mammalian
cells. It helps them to acquire nutrients or other useful substance in order to survive or perform
their functions properly. The entry of molecules into cells is controlled by numerous different
mechanisms depending on the size of molecules. Water, oxygen and glucose can enter cells
simply by diffusion. For other essential small molecules, such as sugars, ions and amino acids,
they can cross the plasma membrane through certain membrane protein pumps, or channels.
However, for macromolecules like proteins or receptors, they must be transported into cells via
membrane-bound vesicles derived from the invagination and pinching-off of pieces of the
plasma membrane in a process called endocytosis(Conner and Schmid, 2003).
Endocytosis (Fig. 4) can take place in multiple ways and are often categorized into two types,
‘phagocytosis’ or cell eating (the uptake of large particles) and ‘pinocytosis’ or cell drinking (the
uptake of fluid and solutes). Phagocytosis usually occurs in specialized cell types, such as
macrophages, monocytes and neutrophils to remove large pathogens or cell debris(Aderem and
Underhill, 1999). Pinocytosis, or fluid phase endocytosis happens in all cells and can be further
divided into four basic subclasses: clathrin-mediated endocytosis (CME), caveolae-mediated
endocytosis, micropinocytosis, and clathrin- and caveolae-independent endocytosis(Conner and
Schmid, 2003; Doherty and McMahon, 2009). In multicellular organism, different endocytic
19
pathways are tightly regulated and working together to control all the complicated physiological
processes and inter- or intra-cellular communications.

Fig. 4. Main pathways of endocytosis. Distinct endocytic pathways can be differed from the size of the
endocytic vesicles (phagocytosis and micropinocytosis have much larger vesical size), their major
molecular composition (clathrin, caveolin or other), the nature of the cargo (ligands, receptors and
lipids) as well as the mechanism of vesical formation.
Reproduced from Conner and Schmid, 2003 with permission of Springer Nature.
20
CHAPTER 3
MATERIALS AND METHODS
3.1 FKBP-ELP gene cloning
ELP gene of A192 was synthesized on pET25b(+) vector (EMD Millipore, Billerica, MA) in
TOP 10 cells (Invitrogen, Carlsbad, CA) by previous members of our laboratory utilizing a
method called recursive directional ligation by plasmid reconstruction(PRe-RDL)(Hassouneh et
al., 2012). FKBP gene was purchased and synthesized on a ampicillin-resistant pIDTsmart vector
(Integrated DNA Technologies, Coralville, IA). Cloning of FKBP-A192 (FA) was performed by
first transferring FKBP gene into a pET25b(+) vector, then digesting the vector and inserting
FKBP gene fragment into another pET25b(+) vector containing the A192 gene. The detailed
procedures were described in a former paper (Dhandhukia et al., 2013). FAF was cloned by
fusing an FKBP gene in frame to the 3’ end of the gene of FA as previously
described(Dhandhukia et al., 2017). The in-frame amino acid sequence of the fusion construct
was confirmed by DNA sequencing.
3.2 FKBP-ELP expression and purification
BLR(DE3) E.coli competent cells (69053, Novagen, Madison, WI) was transformed with
pET25b(+) vector encoding FAF fusion gene. Cells were plated onto agar plates with 100 µg/mL
carbenicillin and incubated in incubator for 16 hours at 37 °C. 6 colonies were picked to select
the highest protein expression yield by inoculating each one in 50 mL autoclaved Terrific Broth
(TB) medium (12105, Mo Bio Laboratories, Carlsbad, CA) supplemented with 100 µg/mL
21
carbenicillin and incubating overnight at 37 °C in an orbital shaker incubator. Cells from each 50
mL starter culture were scaled up by inoculating into 1L TB medium supplemented with 100
µg/mL carbenicillin and allowed to grow for 24 hours at 37 °C in an orbital shaker incubator.
The bacterial culture was then centrifuged at 4000 rpm for 15 mins at room temperature, and the
pellet obtained was re-suspended in phosphate buffered saline (PBS, PBL01, Caisson labs,
Smithfield, UT)). Bacteria in suspension were lysed by sonication for 3 mins with a 10 s on, 20 s
off pulse interval. Polyethyleneimine (PEI, 0.5% w/v final concentration) was added to the cell
lysis to precipitate nucleic acids by centrifugation at 12,000 rpm, 4 °C, 15 mins. The supernatant
which contained fusion ELP proteins was further purified by Inverse Transition Cycling (ITC) as
described before. The colony with highest protein yield was selected and was expressed in 6 L
TB media and purified as described above. The purified protein was filtered using 200 nm sterile
Acrodisc
®
25 mm filters (Pall Corporation, Port Washington, NY). Protein concentration was
calculated by measuring the absorbance at 280 nm on Nanodrop UV-Vis spectrophotometer and
using Beer-Lambert’s Law:
Protein concentration (M) = (A
280
* dilution factor) / (MEC * l) (Equation.1)
Light scattering correction of the A280 value can be made by:
A
280
= A
280
(measured) – 1.929 * A
330
(measured) (Equation.2)
where M is molar concentration, A
280,
A
330
are absorbance at 280 nm and 330 nm respectively, l
is the path length (cm), and MEC is the estimated molar extinction coefficient at 280 nm: 20190
M
-1
cm
-1
for FAF(Pace et al., 1995).
22
3.3 FKBP-ELP physicochemical characterization
The purity and molecular weight of FAF were examined using 4%-20% gradient Tris-Glycine
SDS-PAGE gel (58505, Lonza, Walkersville, MD) by electrophoresis. 5 µg protein in PBS was
mixed with 4X laemmli sample buffer (1610747, Bio-Rad, Hercules, CA) and then loaded onto
the gel. Gel was stained by 10% w/v copper chloride solution and imaged using Bio-Rad
ChemiDoc
TM
Imaging System.
The thermal phase transition behavior of FAF was obtained by measuring its optical density
profile at 350 nm as a function of temperature. A temperature ramp was performed by heating
FAF samples with different concentrations in Beckman Coulter Tm microcells (Brea, CA) on
DU800 UV-Vis spectrophotometer following a 1 °C /min temperature increase and every 0.3 °C
measurement. The maximum first derivative of the optical density at 350 nm was defined as the
phase transition temperature. The transition temperature vs. concentration data was plotted and
fitted into the following linear relationship:
T
t
= b – m [Log
10
(concentration)] (Equation.3)
where b is the intercept temperature (°C) and m is the slope.
3.4 Fluorophore labeling
NHS-Rhodamine (46406, Thermo Fisher Scientific
TM
, Waltham, MA) was dissolved in
anhydrous DMSO (D12345, Invitrogen
TM
, Carlsbad, CA) with a final concentration at 10 mg/mL
according to manufacturer’s protocol. 200 µM FAF was labeled using 2- to 3-fold molar excess
NHS-Rhodamine by reacting at room temperature for 1 hour on a rotator. After 1-hour labeling,
23
Zeba Spin Desalting Column (89893, Thermo Fisher Scientific
TM
, Waltham, MA) was used to
remove the non-reacted dye. The concentration of FAF and rhodamine were measured on
Nanodrop 2000 (Thermo Fisher Scientific
TM
, Waltham, MA) and calculated using equations
below:
Rhodamine concentration (M) = (A
555
* dilution factor) / (MEC * l) (Equation.4)
FAF concentration (M) = (A
280
- CF*A
555
) * dilution factor / (MEC * l) (Equation.5)
where M is molar concentration, A
280,
A
555
are absorbance at 280 nm and 555 nm respectively, l
is the path length (cm) and MEC is the estimated molar extinction coefficient at 280 nm: 20190
M
-1
cm
-1
for FAF, 80000 M
-1
cm
-1
for rhodamine, CF is correction factor which adjusts the
amount of absorbance at 280 nm caused by the dye: 0.34 for rhodamine.
Labeling efficiency is calculated as:
l.e.(%) = C
rho
/ C
pro
* 100% (Equation.6)
The Purity of Rhodamine-FAF (Rho-FAF) was evaluated by 4%-20% gradient Tris-Glycine
SDS-PAGE gel followed by imaging on Bio-Rad ChemiDoc
TM
Imaging System as described
above. Free rhodamine was loaded as positive control.
Notice: The concentration of Rho-FAF mentioned below is usually referred to rhodamine
concentration instead of protein concentration unless otherwise indicated.
3.5 Rapamycin encapsulation and HPLC validation
To encapsulate rapamycin onto Rho-FAF, a two-phase solvent evaporation technique was
utilized. Three times molar excess rapamycin dissolved in a hexane/EtOH mixture (80/20 % v/v)
24
was added to Rho-FAF equilibrated in a glass vial to 37 °C and with protein concentration at
about 200 µM. The two-phase mixture was formed as the organic phase containing rapamycin at
the top and the aqueous phase containing Rho-FAF at the bottom. The organic solvent was
evaporated using mild flow of N
2
with continuously stirring on a magnetic stir plate for 15
minutes and rapamycin was released to be captured by Rho-FAF during this process. The
remaining solution was centrifuged at 13,000 rpm, 37 °C for 10 minutes to remove free drug as
the unbound drug can precipitate into pellets. Centrifugation of the supernatant was repeated for
few times until no pellet was observed. The sample after centrifugation was filtered through 200
nm sterile Acrodisc
®
25 mm filters (Pall Corporation, Port Washington, NY). The protein and
rapamycin concentration of the filtered aliquot was determined using a C-18 RP-HPLC column
(186003033, Waters, Milford, MA) by measuring the absorbance at 280 nm. The HPLC method
was set up and described by previous lab members(Dhandhukia et al., 2017). Concentration was
calculated using two calibrated standard curves respectively.
3.6 Dynamic light scattering
The stability of protein after labeling and encapsulation was evaluated by measuring the particle
size of protein to see if there was an aggregation appeared. Hydrodynamic radius (R
h
) of
Rho-FAF and Rho-FAF/Rapa were determined to represent the particle size by dynamic light
scattering (DLS). 60 µL filtered samples with concentration at 20 µM were loaded in triplicate in
a 384 well black plate (781096, Greiner Bio One, Monroe, NC) and covered with 15 µL mineral
25
oil. The plate was centrifuged at 3,000 rpm for 5 mins to remove air bubbles and was read using
Wyatt Dynapro plate reader (Santa Barbara, CA) at 37 °C.
3.7 Cell culture
MDA-MB-468 cells (HTB-132, ATCC, Manassas, VA) were cultured in Dulbecco’s modified
Eagle’s medium (DMEM)/F-12 medium (DFL21, Caisson labs) supplemented with 10% fetal
bovine serum (FBS, 35-011-CV, Corning, NY) in a humidified incubator with 5% CO
2
at 37 °C.
3.8 Cold competition-binding assay and live cell imaging
To buffer CO
2
change and maintain the medium pH for cells to grow outside of incubator,
complete DMEM/F-12 medium was further supplemented with 25 mM HEPES and filtered for
use in this experiment. MDA-MB-468 cells were seeded on 35 mm glass bottom dishes (MatTek
Corporation, Ashland, MA) at a density of 3*10
5
cells per dish and allowed to grow overnight
for attachment. After around 16-24 hours, culture medium was replaced by 750 µL fresh medium
with HEPES. Cells were placed on ice and pre-cooled Rho-FAF was added to medium at a final
rhodamine concentration of 20 µΜ. After 2-hour incubation on ice, 10-fold excess unlabeled
cold FAF or equal volume of cold Dulbecco’s Phosphate-Buffered Saline (DPBS) were added
and incubated for another 2-hour period on ice. After that, cells were washed three times with
DPBS and 1 mL live cell imaging solution (A14291DJ, Life Technologies, Carlsbad, CA) was
added back to dish with 2 drops of NucBlu
TM
Live ReadyProbes™ Reagent (R37605, Life
Technologies, Carlsbad, CA). Images were captured using the LSM800 confocal microscope
26
(Carl Zeiss Microscopy, Thornwood, NY) mounted on a vibration-free table with a
Plan-Apochromat 63x oil objective.
3.9 Concentration-dependent cellular uptake
To evaluate concentration-dependent intracellular incorporation, cells were trypsinized and
seeded in triplicate into a black bottom 96-well plate at a density of 10,000 cells in 100 µL per
well and allowed to grow overnight to adhere. After around 16-24 hours, culture medium was
replaced with fresh medium follower by treatment with different final concentrations (0, 1, 10,
30, 50, 75, 100 µM) of Rho-FAF. After 16-hour incubation at 37 ºC, medium was aspirated and
cells were washed three times with DPBS. 100 µL live cell imaging solution was added back to
each well and total fluorescence intensity of each well was measured using a plate reader
(BioTek, Winooski, VT).
3.10 Rho-FAF, Fluorescein-dextran and Rhodamine B-dextran cellular uptake
MDA-MB-468 cells were seeded on 35 mm glass bottom dishes at a density of 3*10
5
per dish
and allowed to grow overnight to attach. After around 16-24 hours, culture medium was replaced
by 1 mL fresh medium followed by treatment with 30 µM Rho-FAF and 20 µM
Fluorescein-dextran (70,000 MW, D1823, Life Technologies, Carlsbad, CA), 30 µM Rho-FAF
only, 20 µM Fluorescein-dextran only, 20 µM Rhodamine B-dextran (70,000 MW, D1841, Life
Technologies, Carlsbad, CA) only, and equal volume of DPBS. After 8-hour or 24-hour
incubation at 37 ºC, medium was aspirated and cells were washed three times with DPBS. 1 mL
live cell imaging solution was added back to dish with 2 drops of NucBlu
TM
Live
27
ReadyProbes™. In addition, for the dishes treated with Rho-FAF and Rhodamine B-dextran, 1
µL LysoTracker™ Green DND-26 (L7526, Life Technologies, Carlsbad, CA) was added.
Images were captured using the LSM800 confocal microscope with a Plan-Apochromat 63x oil
objective described as above.
3.11 Time-dependent cellular uptake and loss
To assess the time-dependent uptake, cells were seeded as previously described and treated with
30 µM Rho-FAF or 30 µM Rho-FAF/Rapa for 0h, 1h, 4h, 8h, 16h and 24h at 37 ºC. After
incubation, cells were washed three times with DPBS and imaged in 1 mL live cell imaging
solution with 2 drops of NucBlu
TM
Live ReadyProbes™ using confocal microscope. Integrated
fluorescence intensity and cell number of each image were measured using Fiji (US NIH,
Bethesda, MD).
For the cellular loss experiment, cells were seeded and treated with 30 µM Rho-FAF or 30 µM
Rho-FAF/Rapa for 16 hours pulsed incubation at 37 ºC. After that, culture medium was replaced
with fresh medium and cells were imaged at 0h, 1h, 8h, 24h, 48h and 72h as described above.
Images analysis was also performed as described before. After imaging, cells were washed with
DPBS once and lysed with 120 µL RIPA buffer (89900, Thermo Fisher Scientific
TM
, Waltham,
MA) containing 1.2 µL 100x protease/phosphatase inhibitor (5872, Cell Signaling Technology,
Danvers, MA). Total amount protein of each cell lysate was determined by BCA assay (23227,
Thermo Fisher Scientific
TM
, Waltham, MA) following manufacturer’s protocol. 22.5 µg of total
28
protein was separated on 4%-20% gradient Tris-Glycine SDS-PAGE gel and imaged on Bio-Rad
ChemiDoc
TM
Imaging System.
3.12 Immunofluorescence for mTOR targeting
Cells were grown on glass coverslips in a 12-well plate at a density of 3*10
5
per well and
allowed to attach for overnight followed by treatment with 40 µM Rho-FAF or 40 µM
Rho-FAF/Rapa for 1h, 8h and 16h at 37 ºC. After each time point, cells were fixed with 4%
paraformaldehyde (433689M, Alfa Aesar, Haverhill, MA), permeabilized with 0.1% Triton
X-100 and blocked with 1% bovine serum albumin (BSA, A4503, Sigma-Aldrich, St. Louis, MO)
at room temperature for 1 hour. After that, cells were incubated with a rabbit anti-mTOR
antibody (2983, Cell Signaling Technology, Danvers, MA) for 1 hour at 37 ºC followed by
another hour with Alexa Fluor 488-linked goat anti-rabbit antibody (A32731, Thermo Fisher
Scientific
TM
, Waltham, MA). Cells were then washed and stained with DAPI and mounted on
microscope slides in fluoromount (K048, Diagnostic Biosystems, Pleasanton, CA). Slides were
dried overnight and imaged on a confocal microscope.

29
CHAPTER 4
RESULTS
4.1 Physicochemical characterization of FKBP-A192-FKBP
FAF (Fig. 5a) was successfully expressed and purified from BLR(DE3) E.coli competent cells
by 3-4 rounds of ITC with a yield of 80-90 mg/L. SDS-PAGE was used to determine the purity
and confirm the molecular weight of FAF (Fig. 5b). FAF had a purity around 95% and a
molecular weight about 100 kDa as shown on the gel image. The molecular weight was similar
to both the expected molecular weight based on the amino acid sequence of FAF which is about
97.0 kDa, and the precise molecular weight observed by running samples on MALDI-TOF mass
spectrometer which is about 96.6 kDa(Dhandhukia et al., 2017). There was also a weak band
showing above which was probably the FAF dimer because the protein samples were not mixed
with reducing agent. After drug loading, FAF/Rapa also showed a similar SDS-PAGE profile.
The measurement of optical density change (Fig. 5c) showed that the expressed FAF can
undergo phase separation, confirming that FAF fusion protein retained the thermal responsive
property as observed with ELPs. The transition temperature of FAF was plotted against the
logarithm of FAF concentration, fitted into a linear line using the equation 3, and they were
found in an inverse proportion (Fig. 5d), meaning that higher concentration of FAF displayed a
lower transition temperature. Using the model fitted graph, we can estimate the state of FAF
(soluble or coacervate) at a certain temperature and concentration. At physiological temperature
of 37 °C, FAF should remain as soluble solution within a concentration range of 5-100 µM.
30

4.2 Fluorophore labeling and rapamycin encapsulation
Rhodamine was chosen to label FAF because it is less sensitive to pH and more photostable than
fluorescein. Two- to three-fold molar excess NHS-Rhodamine was used to label the primary

Fig. 5. Physicochemical characterization of FAF. a, A schematic diagram of the structure of
rapamycin binding to FKBP-A192-FKBP fusion protein. FKBP amino acid sequence:
GVQVETISPGDGRTFPKRGQTCVVHYTGMLEDGKKFDSSRDRNKPFKF
MLGKQEVIRGWEEGVAQMSVGQRAKLTISPDYAYGATGHPGIIPPHATLVFDVELLKLE. FAF
amino acid sequence: M-FKBP-G(VPGAG)
192
-FKBP. b, A SDS-PAGE stained by copper chloride
verified the identity and purity of the fusion protein before drug loading and after drug loading. c, The
optical density at 350 nm against temperature profile confirmed the phase transition behavior of the
fusion protein FAF. d, The transition temperature (T
t
) of the fusion protein was found to be
concentration dependent and can be fit into a linear model, T
t
= b – m [Log
10
(concentration)]. b
represents the T
t
at 1 µM which is 63.83 ± 2.1 ºC. m represents the °C change for a 10-fold change in
concentration which is 4.73 ± 1.47 ºC /Log
10
(µM). b, m and figure are shown as Mean ± 95% CI.
31
amino groups (-NH
2
) in FKBP part of the FAF and the release of N-Hydroxysuccinimide
(NHS) facilitated the labeling reaction. The labeling efficiency calculated for two or three-fold
molar excess NHS-rhodamine using equation 4, 5, 6 were 160% and 267% respectively. Zeba
desalting column was used to remove the free rhodamine. 10 µg of protein was loaded onto
SDS-PAGE and the gel showed that no free rhodamine was left in protein samples (Fig. 6a).
A two-phase solvent evaporation technique was used to encapsulate rapamycin onto Rho-FAF
and a C-18 RP-HPLC was utilized to measure the protein and rapamycin concentration. The
protein concentration was about 131 µM and the rapamycin concentration was around 265 µM.

Fig. 6. Rhodamine labeling and rapamycin encapsulation cause no stability issues. FAF was
labelled with NHS-Rhodamine first and rapamycin encapsulation was performed on rhodamine
labelled FAF. a, A SDS-PAGE indicated that FAF was successfully labelled with rhodamine and no
free dye was left after running the desalting column. b, Hydrodynamic radius (R
h
) data showed that
Rho-FAF had a R
h
of 7.8 ± 0.03 nm and Rho-FAF/Rapa had a R
h
of 9.0 ± 0.1 nm at 37 °C (n=3, mean
± SD). There was no significant nanoparticle size change compared to FAF which had a R
h
of 7.9 ±
0.2 nm reported by our group before.
32
The ratio of rapamycin concentration to protein concentration was around 2 to 1 indicating that
the encapsulation was successful.
Dynamic light scattering (DLS) was employed to evaluate the stability of the nanoparticles after
labeling and encapsulation by measuring the hydrodynamic radius (R
h
) of particles. Particles
aggregation is the most common stability issue that leads to an increase in particle size. DLS data
showed that (Fig. 6b) there was no significant nanoparticle size change of Rho-FAF or
Rho-FAF/Rapa compared to FAF which has a R
h
of 7.9 ± 0.2 nm reported by our group
before(Dhandhukia et al., 2017). The rhodamine labeling and rapamycin encapsulation didn’t
cause any aggregation problems and Rho-FAF and Rho-FAF/Rapa should maintain stability like
FAF.
4.3 Cellular uptake of FAF is receptor independent
A cold competition-binding assay using 10-fold excess unlabeled FAF was conducted to
determine the presence of specific cell surface receptor of FAF. The assay was conducted on ice
because at low temperature, uptake process like endocytosis is inhibited due to insufficient
energy but binding is usually not interfered. Cells were first treated with 20 µM Rho-FAF to get
certain amount of binding to cell surface. If there is a selective surface receptor involved in FAF
binding, the addition of 10-fold excess unlabeled FAF should compete off the binding of
Rho-FAF and cause the loss of rhodamine signal. If not, no change of rhodamine signal should
be observed. Interestingly, both images (Fig. 7a) and the value of integrated fluorescence
intensity per cell (Fig. 7b) showed that there was not much difference between the competition
33
group and the PBS control group. Ten-fold excess unlabeled FAF could not compete off the
signal of Rho-FAF, suggesting no specific cell surface receptor was involved upon binding and
thus, the uptake mechanism should not be receptor-mediated endocytosis.

Fig. 7. Ten-fold excess unlabeled FAF cannot compete off the signal of Rho-FAF suggesting no
receptor is involved upon binding. a, MDA-MB-468 cells were pre-incubated with 20 µM Rho-FAF
on ice for 2 hours followed by another 2-hour treatment with either 10-fold excess unlabeled FAF or
equal volume of DPBS on ice. Live cell imaging was performed using confocal microscope. Red:
Rho-FAF; Blue: Hoechst 33342; Scale bar: 20 µm. b, The value of integrated fluorescence intensity
per cell was determined using Fiji (n=2, mean ± SD). There is no significant difference of the value
between 10-fold excess unlabeled FAF competition group and the DPBS control group. 10-fold excess
unlabeled FAF could not compete off the signal of Rho-FAF.
34
4.4 Cellular uptake of FAF is through fluid-phase endocytosis and FAF goes to lysosomes
as the intracellular destination
Since the cellular uptake of FAF was not receptor-dependent and considering the size of FAF,
the most possible mechanism accounting for its uptake should be fluid-phase endocytosis, or
pinocytosis. To determine and verify the endocytosis mechanism of FAF, we used fluorescent
dextran as the fluid-phase endocytosis marker. We co-incubated Fluorescein-dextran and
Rho-FAF with MDA-MB-468 cells to see if they colocalize in cells. We also incubated cells
with Fluorescein-dextran alone as a negative control. Images were taken using confocal
microscope and Mander’s colocalization coefficient (MCC)(Manders et al., 1993) was calculated
for both red (Rho-FAF) channel and green (Fluorescein-dextran) channel using ZEN2009
software (Carl Zeiss Microscopy, Thornwood, NY). MCC was measured for each channel
separately by adding the pixels in the colocalized region and then dividing by the sum of pixels
in each channel. Each pixel has a value of 1 and MCC values range from 0 to 1. The equation for
calculating the MCC for Red (R) and Green (G) channel are shown as below:
𝑀
"#$
=
&
',)*+*) '
&
' '
, 𝑀
,"##-
=
.
',)*+*) '
.
' '
(Equation.7)
The colocalization coefficient for Rho-FAF and Fluorescein-dextran are 0.77 ± 0.12 and 0.76 ±
0.11 respectively (Fig. 8b), suggesting that they two were highly colocalized with each other. By
visually identifying in images (Fig. 8a), Rho-FAF and Fluorescein-dextran also followed very
similar distribution pattern in cells and there were obvious yellow appearing in the region of
overlap parts, further supporting that FAF has a similar uptake mechanism and intracellular
35

Fig. 8. Rho-FAF colocalizes with Fluorescein-dextran upon cellular uptake and the
internalization of Rho-FAF is not saturable. a, MDA-MB-468 cells were co-incubated with 30 µM
Rho-FAF and 20 µM Fluorescein-dextran for 8 hours and live cell imaging was conducted by confocal
microscopy. Images showed highly colocalization between red channel and green channel, indicating by
the appearance of yellow color. Red: Rho-FAF; Green: Fluorescein-dextran; Blue: Hoechst 33342. b,
Colocalization coefficient of Rho-FAF (Avg.=0.77) and Fluorescein-dextran (Avg.=0.76) of each
individual cell were measured using ZEN2009 software as described before, confirming the highly
colocalization between FAF and dextran (n=16, Mean ± SD). c, MDA-MB-468 cells were treated with
a serial of Rho-FAF with increasing concentrations for 16 hours and total fluorescence intensity was
measured by a plate reader. The internalization of Rho-FAF was increasing as concentration
increased, showing no saturation in terms of cellular uptake (R
2
=0.89, n=4-6, Mean ± SD).
36
trafficking pathway as dextran.
To further proved that the cellular uptake of FAF was through fluid-phase endocytosis and no
receptor was involved, we performed a cell uptake assay to examine the saturability of
internalization when cells were treated with a serial of Rho-FAF with increasing concentrations
(1-100 µM). When the internalization of cargoes is mediated by specific receptors, it usually
causes an uptake saturation due to the limited amounts of receptors. However, when the uptake
of cargoes is mediated by pinocytosis and no receptor is involved in internalization, cells should
incorporate cargoes by engulfing the extracellular fluid containing cargoes around it. In this case,
no saturation should be observed. The total fluorescence intensity was measured to represent the
amount of internalization and was plotted against concentration (Fig. 8c). The figure showed that
the internalization of Rho-FAF increased as concentration went up. A linear model was used to
fit the graph with a R
2
about 0.89. The good fit into linear line suggested that the cellular uptake
of Rho-FAF was not saturable within that concentration range. This observation further
confirmed that FAF was internalized through fluid phase endocytosis without specific cell
surface receptor.
Furthermore, the final intracellular fate of FAF was explored, specifically trying to determine
whether FAF will go to lysosomes since lysosomes are usually considered as the final destination
of endocytic uptake and as the place for cargo releasing and biodegradation. Because we
observed before that FAF was highly colocalized with dextran upon uptake, we also incubated
cells with Rhodamine-B dextran to see where dextran will go. After incubation with either
37

Fig. 9. Rho-FAF and Rho B-dextran show the same colocalization pattern with lysosomes. a,
MDA-MB-468 cells were either incubated with 30 µM Rho-FAF or 20 µM Rhodamine B-dextran for
8 hours followed by addition of 1 µL LysoTracker green before imaging. Live cell imaging was
conducted by confocal microscopy and colocalization coefficient was measured using ZEN2009
software. Images showed highly colocalization between red channel and green channel, indicating by
the appearance of yellow color. Red: Rho-FAF or Rho B-dextran; Green: Fluorescein-dextran; Blue:
Hoechst 33342. b, Colocalization coefficient revealed that FAF and dextran shared the same
colocalized pattern with lysosomes with a large portion of red signal colocalizing with a small portion
of green signal (n=12, 22, Mean ± SD).
38
Rho-FAF or Rho B-dextran, lysotracker green was used to mark lysosomes and live cell imaging
was performed on confocal microscope. Confocal images showed that both FAF and dextran
displayed certain amounts of colocalization with lysosomes (Fig. 9a). Mander’s colocalization
coefficient for both red and green channel were measured, showing a very similar colocalization
pattern of FAF and dextran with lysosomes (Fig. 9b). The MCC for Rho-FAF and LTG were
0.78 ± 0.22 and 0.24 ± 0.13 (n=12, Mean ± SD) respectively, while the MCC for Rho B-dextran
and LTG were 0.85 ± 0.10 and 0.31 ± 0.10 (n=22, Mean ± SD) respectively. The colocalization
coefficient for either FAF or dextran was much higher than the one for lysotracker green,
meaning that a large portion of the red population was colocalized with a much smaller portion
of the green population. In other words, Rho-FAF and Rho B-dextran were colocalized with
lysosomes but only with a small fraction of all the lysosomes in MDA-MB-468 cells. This
experiment indirectly confirmed that the intracellular fate for FAF and dextran were similar. FAF
followed the same pattern as dextran after being internalized into MDA-MB-468 cells and both
went to lysosomes as their final fates.
4.5 Cellular uptake and loss of Rho-FAF and Rho-FAF/Rapa are time dependent.
Except concentration, time is another important factor that can impact cellular uptake efficiency.
The time needed for the incorporation and degradation of FAF in cells is crucial to evaluate its in
vivo application. We performed both continuous incubation and pulsed incubation assays with
both Rho-FAF and Rho-FAF/Rapa on MDA-MB-468 cells to collect the time-dependent kinetic
profiles of uptake and loss respectively. In the continuous incubation assay, kinetic cellular
39
uptake was assessed. MDA-MB-468 cells were incubated with 30 uM rhodamine concentration
materials for different time intervals. Confocal images (Fig. 10a) showed that increased
fluorescence signal was observed in longer incubation time points. The value of integrated
fluorescence intensity per cell was quantified and plot it against time (Fig. 10b). Both

Fig. 10. Time-dependent internalization profile of Rho-FAF with or without rapamycin. a,
MDA-MB-468 cells were incubated with 30 µM Rho-FAF (red) for 1h, 4h, 8h, 16h and 24h at 37 °C
and imaged using confocal microscopy. Increased fluorescence signal was observed in longer
incubation time points. b, At least 3 images of each time point were took and analyzed for
fluorescence intensity and cell number. Fluorescence intensity/cell against time profile was plotted
showing the time-dependent uptake (n=3-7, Mean ± SD).
40
images and the plot indicated no difference between Rho-FAF and Rho-FAF/Rapa and no
saturation trend within 24 hours was observed for both. We expect that ultimately, a plateau will
occur after 24 hours where the entry and excretion of Rho-FAF or Rho-FAF/Rapa reach a
balance.
In the pulsed incubation assay, kinetic cellular degradation was evaluated. Cells were pulsed
treated with Rho-FAF and Rho-FAF/Rapa for 16 hours and subsequently imaged and lysed after
different incubation time intervals. Decreased fluorescence signal was observed in longer
incubation time point, indicating Rho-FAF and Rho-FAF/Rapa were being degraded (Fig. 11a).
The plot of integrated fluorescence intensity per cell against time was fitted into a one-phase
decay model (Fig. 11b) showing the cellular loss half-life of Rho-FAF or Rho-FAF/Rapa were
17.73 hours and 21.27 hours respectively. Cell lysates were resolved on a SDS-PAGE and
fluorescent gel was captured (Fig. 11c) showing that the main FAF band was diminishing as time
increased with multiple smears below appeared, further confirming that Rho-FAF was being
degraded in MDA-MB-468 cells. Higher value of integrated fluorescence intensity per cell were
observed in Rho-FAF/Rapa group, which probably related to the anti-proliferative property of
rapamycin that can inhibit cell growth. In the Rho-FAF group without rapamycin, cells were kept
dividing so Rho-FAF would get diluted because the number of cells increased. This drop of
integrated fluorescence intensity per cell value was not because of degradation but cell division.
However, based on the SDS-PAGE image, the intensity of band had at least 10-fold loss at 72h
time point compared to 0h, which is a much higher fold loss than if the loss of intensity only
41
caused by cell division (the number of cells can increase at most 2-2.5 fold for 72 hours). Thus,
the effect of cell division should only account for a minimal part of the decreased fluorescence

Fig. 11. Cellular loss of Rho-FAF and Rho-FAF/Rapa. a, MDA-MB-468 cells were pulsed
incubated with 30 µM Rho-FAF or Rho-FAF/Rapa (red) for 16 hours at 37 °C. After changing
medium, cells were imaged at 0h, 1h, 8h, 24h, 48h and 72h using confocal microscopy. Decreased
fluorescence signal was observed in longer incubation time point, indicating ELPs were being
degraded. b, Fluorescence intensity/Cell was measured and calculated as described previously. The
plot (n=3-7, Mean ± SD) was fitted into a one-phase decay model showing the cellular loss half-life of
Rho-FAF or Rho-FAF/Rapa were 17.73 hours (15.55-20.38 hours, 95% CI) and 21.27 hours
(17.65-26.52 hours, 95% CI) respectively. c, A fluorescent SDS-PAGE showed the main band for
FAF was diminishing as time increased with multiple bands below appeared. Estimated half-life for
Rho-FAF using densitometry was 18.98 hours (15.35-21.59 hours, 95% CI), which is similar to the
one estimated by confocal images.
42
signal while most of it should be explained by degradation. Some chemical reactions like
oxidation on the rhodamine in cells can also be one of the reasons for the fluorescence signal loss
which details remained unknown. If an immunoblot can be performed on those cell lysates using
anti-ELP or anti-FKBP antibody to confirm that not only the fluorescence signal gets decreased
but also the amount of protein becomes less, it would be a stronger proof that the loss of
fluorescence signal is due to the protein degradation.
4.6 Rho-FAF/Rapa targeting mTOR in MDA-MB-468 cells
Since mTOR is the molecular target for FKBP-Rapamycin complex, we performed
immunofluorescence assays to investigate whether Rho-FAF/Rapa will target mTOR in
MDA-MB-468 cells. Cells were treated with Rho-FAF/Rapa for 1h, 8h and 16h and
subsequently fixed, permeabilized and stained with anti-mTOR antibody. Not much
colocalization between Rho-FAF/RAPA and mTOR was observed at 1h time point, but in 8h and
16h time point, certain colocalization was seen as indicated by the white arrows (Fig. 12a).
However, the percent of cells that had colocalization (around 20%) and the colocalization
coefficient for Rho-FAF/Rapa (around 0.3-0.5) were low. Another immunofluorescence assay
was conducted to confirm that mTOR and lysosomes were associated with each other using
anti-mTOR antibody and anti-LAMP1 antibody (Fig. 12b). There were obvious white colors in
the overlap regions indicating that mTOR was localized to the lysosomes under activated state in
MDA-MB-468 cells. In both immunofluorescence assays, only small parts of mTOR population
showed colocalization with either Rho-FAF/Rapa or lysosomes. One of the reasons is that the
43

Fig. 12. Rho-FAF/Rapa targeting mTOR and mTOR localization in MDA-MB-468 cells. a,
MDA-MB-468 cells were treated with 30 µM Rho-FAF/Rapa (red) for 1h, 8h and 16h at 37 °C. After
incubation, cells were fixed, permeabilized and mTOR (green) was stained using an anti-mTOR
antibody followed by an alexa fluor 488-linked anti-rabbit secondary antibody. Certain amount of
colocalization between mTOR and FAF was observed in 8h and 16h time point, whereas very less
was seen in 1h time point. b, Immunofluorescence was performed using anti-mTOR antibody and
anti-LAMP1 antibody as primary antibodies to detect mTOR and lysosome localization in
MDA-MB-468 cells. Overlay image showed that mTOR and lysosome were associated together in
cells under nutritive state.
44
anti-mTOR antibody used in the experiments can detect both mTORC1 and mTORC2 but only
mTORC1 was supposed to colocalize with Rho-FAF/Rapa or lysosomes.
3.7 Cellular internalization mechanism of FAF or FAF/Rapa
Based on all the experimental results, we proposed a schematic illustration (Fig. 13) of the
cellular uptake mechanism of MDA-MB-468 cells after treatment with FAF or FAF/Rapa
nanoparticles. FAF/Rapa was internalized into cells via pinocytosis without specific cell surface
receptor involved. FAF/Rapa then moved to lysosomes as their intracellular destination and was
degraded and released to bind to its final target, mTORC1 which was associated with lysosomes
under nutrient-rich condition to trigger downstream signaling pathway and performed its
anticancer function.

Fig. 13. Schematic of FAF/Rapa on cellular uptake pathways in MDA-MB-468 cells.
45
CHAPTER 5
DISCUSSION
Very limited papers reported the cellular uptake mechanism of ELPs. One group fused ELPs with
CPP to facilitate the delivery of a therapeutic peptide and they revealed that the internalization of
ELPs and CPP-ELPs were through caveolae-independent endocytic pathways(Bidwell and
Raucher, 2010). However, another group utilized supercharged ELPs to enhance the cellular
uptake of GFP and found that GFP-ELPs was mainly incorporated via caveolae-mediated
endocytosis(Pesce et al., 2013). Thus, it is hard to predict and compare the cellular uptake
mechanism for different ELPs or ELP fusion proteins since they all have distinct components and
properties. In the present study, FAF was only found to be internalized through fluid phase
endocytosis as it highly colocalized with the fluid phase endocytosis marker, dextran and its
cellular uptake had no saturation within 1-100 µM range (Fig. 8). However, as mentioned before,
there are four different pinocytosis pathways and which specific one is dominated in FAF uptake
remained unknown. Usually, molecules can be internalized through one particular pathway, but
there might be multiple pathways involved. To better understand that, we can use approaches to
block one of those pathways like using non-specific chemical inhibitors, selective pharmacologic
agents or genetic methods and see if that will change the uptake of molecules(Dutta and
Donaldson, 2012).
FAF went to lysosomes as their intracellular destination (Fig. 9) and FAF and FAF/Rapa were
degraded in MDA-MB-468 cells as less rhodamine signal was seen in longer incubation time
46
points (Fig. 11). Although materials endocytosed by fluid phase endocytosis are typically
destined to lysosomes for degradation(Baravalle et al., 2005), whether FAF is degraded in
lysosomes or not still remained unclear. In the future, we can perform the cell uptake pulsed
incubation assay with lysosomal protease inhibitors like chloroquine, ammonia or
leupeptin(Seglen et al., 1979) to see if that can stop the degradation. Another question regarding
the degradation is that which parts of the construct are being degraded. We think most probably,
it is the ELPs parts that are subjected to digest. An immunoblot using anti-ELPs antibody can be
conducted on SDS-PAGE in Fig. 11c to verify that the following smear bands are ELPs.
Alternatively, we can also use anti-FKBP12 antibody. The rationale is that the endogenous
FKBP12 should be around 12 kDa and if anti-FKBP12 antibody can detect any bands above that,
it means that FKBP12 is still linked to ELPs and it is the degradation of ELPs that leads to
different molecular weights showing as multiple bands.
Immunofluorescence studies were able to provide some proofs for FAF/Rapa targeting mTOR in
MDA-MB-468 cells (Fig. 12a). However, interestingly, we also saw some localizations between
FAF and mTOR when we only incubated cells with FAF alone since we knew that without
rapamycin, FKBP cannot bind to mTOR. One possible reason can be FAF was actually
associated with lysosomes and since lysosomes and mTOR were near to each other, confocal
microscope may not have enough resolution to distinguish. Thus, we still need more solid
evidence to support mTOR, lysosomes, FAF/Rapa association may be by techniques like
immunoprecipitation since microscopy and images have their limitations.
47
Although in general, this study provides some useful information about FAF uptake mechanism,
one important part is missing here which is the evidence of drug activity. The evidence of drug
activity can be accomplished by performing the p70S6 kinase immunoblot of the cell lysates
from the cells which are treated with FAF/Rapa in the presence and absence of endocytosis
inhibitors and lysosomal protease inhibitors. If we observe the reversal of activity without uptake
and degradation, it can also prove that the active drug remains bound to the carrier upon cellular
uptake.

48
CHAPTER 6
CONCLUSION
In conclusion, we successfully expressed and purified FAF fusion protein and performed
rhodamine labeling and rapamycin encapsulation on it. In MDA-MB-468 cells, FAF is
internalized through fluid phase endocytosis without specific cell surface receptor involved. It is
biodegradable and goes to lysosomes as its intracellular destination. FAF/Rapa associates with
mTOR but stronger evidence need to be presented. Overall, we provide a basis for tracking
endocytosed ELPs, but more studies need to be carried out to fully understand the whole picture of
their intracellular fates.

49
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Creator Pan, Xiaoli (author)

Core Title Cellular uptake mechanism of elastin-like polypeptide fusion proteins

School School of Pharmacy

Degree Master of Science

Degree Program Molecular Pharmacology and Toxicology

Publication Date 08/02/2019

Defense Date 05/16/2018

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag cellular uptake,confocal microscopy,drug delivery,elastin-like polypeptides,FK506 binding protein,OAI-PMH Harvest,rapamycin

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Advisor Mackay, John Andrew (committee chair), Duncan, Roger (committee member), Okamoto, Curtis (committee member)

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