University of Connecticut University of Connecticut

OpenCommons@UConn OpenCommons@UConn

University Scholar Projects University Scholar Program

Spring 5-5-2019

Development of a Sonically Powered Biodegradable Development of a Sonically Powered Biodegradable

Nanogenerator for Bone Regeneration Nanogenerator for Bone Regeneration

Avi S. Patel avipatel97@gmail.com

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Recommended Citation Recommended Citation Patel, Avi S., "Development of a Sonically Powered Biodegradable Nanogenerator for Bone Regeneration" (2019). University Scholar Projects. 64. https://opencommons.uconn.edu/usp_projects/64

Patel 1

Development of a Sonically Powered

Biodegradable Nanogenerator for Bone

Regeneration

Cover Page Avi S. Patel

University Scholar

BA. Individualized: Health, Medicine, and Society

BS. Molecular and Cell Biology

University of Connecticut

05 May 2019

Thanh Duc Nguyen, Ph.D.

Principal Investigator, University Scholar Primary Advisor, MCB Thesis Advisor

David Goldhamer, Ph.D.

MCB Honors Advisor & University Scholar Advisor

Maryann Morris, MS, RD, CD-N

University Scholar Advisor & Individualized Degree Advisor

Kathryn Ratcliff, Ph.D.

University Scholar Advisor & Individualized Degree Advisor

Pamela Erickson, Ph.D.

Individualized Degree Primary Advisor

XiaoBin Huang, Ph.D.

Post-Doctoral Research Assistant

Ritopa Das & Eli Curry

Graduate Research Assistants

Patel 2

Abstract

Background: Reconstruction of bone fractures and defects remains a big challenge in orthopedic surgery.

While regenerative engineering has advanced the field greatly using a combination of biomaterial

scaffolds and stem cells, one matter of difficulty is inducing osteogenesis in these cells. Recent works have

shown electricity’s ability to promote osteogenesis in stem cell lines when seeded in bone scaffolds;

however, typical electrical stimulators are either (a) externally housed and require overcomplex

percutaneous wires be connected to the implanted scaffold or (b) implanted non-degradable devices

which contain toxic batteries and require invasive removal surgeries.

Objective: Here, we establish a biodegradable, piezoelectric Poly-L-Lactic Acid (PLLA) scaffold that uses

external, non-invasive ultrasound to generate an electric charge that promotes stem cell osteogenesis.

Methods: Demonstration of this system included (1) development of a piezoelectric PLLA mesh, (2)

verification of its piezoelectric efficacy and degradation, (3) manufacturing of a PLLA scaffold, (4) in vitro

testing of the system’s ability to enhance bone regeneration compared to a control, and (5) using

assessments of cell proliferation and differentiation through protein, mineral, and gene assays.

Results: Ultimately a 3000rpm electrospun PLLA nanofiber film that could output 40mV when stimulated

with 40kHz 0.4W/cm2 ultrasound was assembled into a bone scaffold and seeded with adipose-derived

stem cells (ADSCs). In vitro testing showed that relative to a control, in cells subjected to the experimental

conditions alkaline phosphatase production increased 5-fold, mineral production increased 18-fold,

osteocalcin gene 40-fold, and osterix gene 100-fold.

Conclusion: The production of surface-level charge from ultrasonic stimulation of PLLA and the use of that

charge to promote osteogenic differentiation in ADSCs was successfully demonstrated. The fact that PLLA

was successfully used in combination with externally applied ultrasound to produce electrical charge

opens up new frontiers for the field of tissue regeneration. This advancement helps make tissue

engineering a tool that can tackle problems of even greater magnitude.

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Acknowledgments

Thank you to Dr. Thanh Nguyen for the incredible opportunity he gave me four years ago to become an

undergraduate research in his lab and for all the help he has provided to help me achieve my goals. Thank

you to Dr. David Goldhamer for supervising my project and for whose critiques helped shape this paper.

Thank you to Ms. Maryann Morris whose constant support of my ambitions gave me the confidence to

pursue a broader degree related to public health. Thank you to Dr. Kathryn Ratcliff, who introduced me

to the social determinants of health, a fundamental aspect of healthcare that I would have otherwise

overlooked. Thank you to Dr. Pamela Erickson, whose guidance and instruction influenced me to take such

influential classes in medical anthropology and cultural anthropology that greatly shaped my interests in

medicine. Thank you to XiaoBin Huang, Ritopa Das, and Eli Curry, for introducing me to the world of

research, for letting me work alongside them, and teaching me many lessons along the way. Thank you

also to the University Scholars Program, the Honors Program, and the STEM Scholars program for helping

fund my plan of study. Lastly, but most of all, thank you to all my friends here at UCONN for believing in

me, respecting me, and loving me every step of the way.

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Table of Contents

Cover Page .................................................................................................................................................... 1

Abstract ......................................................................................................................................................... 2

Acknowledgments ......................................................................................................................................... 3

Table of Contents .......................................................................................................................................... 4

List of Figures ................................................................................................................................................ 5

Keywords ....................................................................................................................................................... 5

List of Abbreviations & Acronyms ................................................................................................................. 6

Introduction .................................................................................................................................................. 7

Introduction to Tissue Engineering ........................................................................................................... 7

Introduction to Bone Regeneration .......................................................................................................... 7

Introduction to Use of Physical Stimulation in Bone Regeneration ......................................................... 8

Mechanical Forces .............................................................................................................................. 10

Ultrasound .......................................................................................................................................... 11

Shock Waves ....................................................................................................................................... 12

Laser .................................................................................................................................................... 13

Introduction to Use of Electrical Physical Stimulation ............................................................................ 14

Capacitive Coupling ............................................................................................................................. 15

Inductive Coupling .............................................................................................................................. 16

Direct Current ..................................................................................................................................... 17

Introduction to Direct Current Surface Charge Electrical Stimulator Systems ....................................... 17

Introduction to Use of Piezoelectrics for Surface Charge Electrical Stimulators .................................... 18

Introduction to Use of Poly-L-Lactic Acid for Surface Charge Electrical Stimulators .............................. 18

Materials & Methods .................................................................................................................................. 20

Step 1. Processing and Manufacturing of the Piezoelectric PLLA Nanofiber Mesh................................ 20

Step 2. Verification of PLLA Nanofiber Mesh Efficacy and Degradation ................................................ 22

Step 3. Assembly of the PLLA Scaffold .................................................................................................... 23

Step 4. Test of Osteogenic Differentiation in vitro ................................................................................. 23

Step 5. Assessment of Cell Proliferation and Differentiation ................................................................. 26

Results ......................................................................................................................................................... 29

Step 1. Processing and Manufacturing of the Piezoelectric PLLA Nanofiber Mesh................................ 29

Step 2 & 3. Verification of PLLA Nanofiber Mesh Efficacy and Degradation & Assembly of the PLLA

Scaffold ................................................................................................................................................... 30

Step 4 & 5. Test of Osteogenic Differentiation in vitro & Assessment of Cell Proliferation and

Differentiation ......................................................................................................................................... 31

Discussion.................................................................................................................................................... 33

Conclusion ................................................................................................................................................... 38

References .................................................................................................................................................. 39

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List of Figures

Figure 1. Schematic of Various Physical Stimulation Enhancements For Bone Regeneration ..................... 9

Figure 2. Main Methods of Administering Electric Stimulation to Bone .................................................... 15

Figure 3. Piezoelectric PLLA Nanofiber Mesh Manufacturing Diagram ...................................................... 20

Figure 4. Diagram of the Completed PLLA Bone Graft ............................................................................... 23

Figure 5. Table of Experimental Groups ..................................................................................................... 24

Figure 6. Diagram of Ultrasound Stimulation Setup ................................................................................... 25

Figure 7. SEM Images of Non-Piezoelectric PLLA Nanofiber Film ............................................................... 29

Figure 8. SEM Images of Piezoelectric PLLA Nanofibers ............................................................................. 29

Figure 9. Verification of PLLA Piezoelectric Output .................................................................................... 30

Figure 10. Graph of PLLA Degradation Study .............................................................................................. 30

Figure 11. ADSC Protein and Mineral Expression Test Results ................................................................... 31

Figure 12. ADSC Gene Expression Test Results ........................................................................................... 31

Figure 13. BMDSC Reporter Cell Activity .................................................................................................... 32

Keywords

Biodegradable, Bone Regeneration, Piezoelectric, Poly-L-Lactic Acid (PLLA), Ultrasound

Patel 6

List of Abbreviations & Acronyms

adenosine triphosphate (ATP), 13

adipose-derived stem cells (ADSCs), 8

adipose-derived mesenchymal stem cells

(ADSCs), 10

alkaline phosphatase (ALP), 13

analysis of variance (ANOVA), 27, 28

barium titanate (BaTiO3), 18

bicinchoninic acid (BCA), 26

bone marrow-derived stem cells (BMDSCs), 8

bone morphogenetic proteins (BMPs), 13

bone sialoprotein (BSP), 11

calcium (Ca2+), 11

Capacitive coupling (CC), 15

collagen type I (COL I), 14

dentin matrix protein (DMP1, 13

dichloromethane (DCM), 20

differential scanning calorimetry (DSC), 22

Dimethylformamide (DMF), 20

Direct current (DC), 15

electric field (EF), 15

electrical stimulation (ES), 14

extracellular matrix (ECM), 8

extracorporeal shock wave therapy (ESWT), 12

extremely low-frequency pulsed

electromagnetic field (ELF-PEMF), 16

Food and Drug Administration (FDA), 11

glycosaminoglycans (GAGs), 15

Honest Significant Difference (HSD), 27

Inductive coupling (IC), 15

lead zirconate titanate (PZT), 18

low-intensity pulsed ultrasound stimulation

(LIPUS), 11

Low-level laser therapy (LLLT), 13

mCherry red fluorescent protein reporter for

dentin matrix protein (DMP1-RFP-mCherry),

28

mesenchymal stem cells (MSCs), 10

osteoarthritis (OA), 13

osteocalcin (OCN), 11

osteopontin (OPN), 11

phosphate buffer solution (PBS), 22

p-nitrophenyl phosphate (pNPP), 26

polydimethylsiloxane (PDMS), 22

poly-L-lactic acid (PLLA), 18

polyvinylidene fluoride (PVDF), 18

pulsed electromagnetic fields (PEMFs), 15

quantitative polymerase chain reaction (qPCR),

26

runt-related transcription factor 2 (RUNX2), 11

scanning electron microscopy (SEM), 21

topaz green fluorescent protein reporter for

bone sialoprotein (BSP-GFP-topaz), 28

transforming growth factor-beta 1 (TGF-β1), 16

ultrasound (US), 11

X-ray diffraction (XRD), 22

zinc oxide (ZnO), 18

Patel 7

Introduction

Introduction to Tissue Engineering Classical tissue regenerative engineering is an interdisciplinary field of advanced material science,

cell biology, and developmental biology, with the aim of promoting the regeneration of complex tissues

and organs [1]. In this process, natural or synthetic scaffolds, cells, and growth factors combine to form a

construct, structurally, functionally, and mechanically similar to the native tissue that requires repair [2].

The field began around the 1970s when chondrocytes were for the first time seeded onto spicules of bone

[3]. Although no new tissue was generated, it revealed the importance of the substrate cells were seeded

on. Whereas naturally occurring scaffolds having physical and chemical properties similar to that of the

native tissue, they could not be manipulated for experimentation or improvement thus resulting in

unpredictable outcomes. When this idea developed in the 1980s into the concept of designing artificial

scaffoldings for cell delivery as opposed to seeding cells onto available natural substrates, the true birth

of tissue engineering occurred – one focused less on the cells themselves and more on how they can be

physically and chemically directed towards growth [3].

Introduction to Bone Regeneration It is well known that bone disorders such as osteoporosis, osteoarthritis, and bone fractures,

commonly occur due to abnormal physiology or physical injury. Reconstruction of bone fractures and

defects has for a long time been an area of focus in orthopedic surgery [4]. The most common treatment

options, replacement allografts and autografts, face many issues. Allografts, bone grafts from a donor,

usually suffer from problems of limited supply, donor site infection, and host immune rejection [5].

Autografts, grafts sourced from other parts of that individual’s body invite additional complications at the

donor site [5]. The second most common treatment option, long bone fixation, is highly invasive, risky,

and painful [4]. Regenerative and tissue engineering strategies, which employ a synergistic combination

of biomaterial scaffolds, stem cells, and/or small-molecule therapies, have therefore emerged as an

important area of study [2].

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Bone regenerative engineering specifically aims to create stable, bioactive, and native tissue-like

scaffolds that can repair bone damages [6]. These scaffolds are often combined with osteogenic or stem

cells to create replacement tissue grafts with enhanced regenerative capability. Several techniques and

strategies have emerged to promote bone regeneration. However, many of these novel treatments still

suffer from issues of their own. One great challenge and key matter of difficulty is inducing osteogenesis,

bone formation, in adipose-derived stem cells (ADSCs) and bone marrow-derived stem cells (BMDSCs) and

reconstructing tissues with sufficient mechanical strength and native tissue-like function [7].

When this differentiation occurs erroneously, fibrous connective tissue rapidly occupies the bony

defect rather than normal bone formation occurring [8]. The resulting fibrous connective tissue buildup,

with its low mechanical strength and cartilage-like structure, creates defective bone [8]. To address these

issues, people have researched biochemical stimuli including enhanced blood plasma, novel biomaterial

scaffolds, and various growth factors that can push stem cells away from the fibrous connective tissue

direction of differentiation and towards the bone side of regeneration; however, most attention was put

into the chemical and biological behaviors [9]–[12]. Physical stimulation of bone has been not extensively

researched or utilized clinically in bone regeneration efforts and warrants a greater understanding.

Introduction to Use of Physical Stimulation in Bone Regeneration Since bone is exposed to multiple internal and external physical forces, however, biomechanical

environment plays an important role in maintaining, repairing, and remodeling its tissues to meet

functional demands and maintain the tissue homeostasis [13]. In fact, the physical properties of the cell

microenvironment are equally important as the biochemical properties. For example, it has been shown

that altering the stiffness of the extracellular matrix (ECM) could direct stem cell differentiation, with

increasing stiffness directing differentiation toward more mechanically competent tissues, such as

cartilage and bone, and away from the more delicate adipose and neuronal tissues [12]. Physical stimuli,

including mechanical forces, ultrasound, shock waves, laser, electromagnetism, and electrical have

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already shown active roles in bone regeneration and fracture healing in vitro and in vivo [14]. Figure 1

summarizes these common physical stimulations.

All of these physical stimulations have significant impacts on cell fate and behavior through

various intracellular signaling pathways. This suggests that the use of such stimuli can be a promising

strategy to improve bone fracture healing and regeneration. To date, some physical manipulations have

already been introduced into clinical applications for bone regeneration, however, others have faced

some difficulties. Knowledge of these advancements in physical stimulation for bone regeneration is

important for the development of future smart tissue grafts that can repair bone in novel ways.

Figure 1. Schematic of Various Physical Stimulation Enhancements For Bone Regeneration: (1) Step one of all tissue engineering strategies is harvesting the appropriate stem cell line. (2) Step two is seeding the cells in an artificial scaffold. In this case, the scaffold used is a piezoelectric PLLA scaffold. (3) Step three in this method of bone regeneration is the application of physical stimulation. The four most common types are mechanical, ultrasound, shock wave, and electromagnetic stimulation. (4) The last step that is also interchangeable with step three is implanting the scaffold and continuing the stimulation.

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Mechanical Forces Mechanical forces are one type of physical stimulation that has been adopted into clinical practice

[15]. It is well known that both extrinsic and intrinsic mechanical forces can induce tissue resistance and

adaptation. The induced tissue forces are transmitted to the micromechanical environment of resident

cells and thus influence the intracellular forces. Cells can subsequently modify their micromechanical

environments via cytoskeletal rearrangement or molecular cascade transduction activation. This

ultimately alters the synthesis or degradation of the extracellular matrix and feeds back to alter cellular

sensitivity to incoming mechanical forces [16]. Studies have demonstrated that appropriate mechanical

forces are important for normal bone cell localization, orientation, metabolism, and homeostasis [17]. The

two most common mechanical forces that contribute to this are cyclic strain and fluid shear stress [18].

Cyclic strain describes the cycles of loading and unloading that cause the compression and

relaxation of the ECM, which induce strain on the bone cells. Cyclic strain includes repeated tensile strain

(pulling apart) as well as compressive strain (pushing together) of the tissue. Bone is constantly exposed

to cyclic strain when an individual body is moving in daily life. The magnitude of tensile strain is important

in bone development and the fate determination of mesenchymal stem cells (MSCs) and has reportedly

been related to inhibition of adipogenesis (a process balancing osteogenesis and chondrogenesis) [19]. In

mouse adipose-derived mesenchymal stem cells (ADSCs), cyclic tensile strain has also been shown to

significantly reduce adipogenesis [20]. Studies have shown cyclic strain could increase the bone-to-

adipose ratio by upregulating the expression of actin-associated proteins, and activation of stretch-

activated cation channels [21]. Elements of the cytoskeleton bridging actin fibers to the nuclear

membrane also have been shown to play an important role in osteogenesis during this mechanical

stimulation [22].

The other common mechanical force that has been studied to aid in bone development and stem

cell osteogenesis particularly is fluid shear stress -- the pulsatile or oscillating shear stress on the

Patel 11

musculoskeletal system caused by the flow of blood in the circulatory system [23]. Studies have proved

the application of both continuous flow and pulsating fluid flow increases osteogenic differentiation of

ADSCs as compared to static cultures [24]. Studies of fluid shear stress have shown pulsating fluid flow to

produce the greater osteogenic induction with gene expression of runt-related transcription factor 2

(RUNX2), an indicator of early osteogenesis, increasing significantly and expression of osteopontin (OPN),

an indicator of late osteogenesis, remaining unchanging after 3 hours of application [25]. These studies

suggest pulsating fluid flow may affect the early stages, but not the late stages of osteogenic

differentiation [25]. The enhancement of osteogenesis from fluid flow relates to the distribution of

nutrient and growth factors in the cell with enhanced expression of bone-specific markers being found in

a uniform distribution in perfusion cultures and only the outer regions in static cultures [24]. Thus, the

improved osteogenesis from the fluid flow may be attributed to the better distribution of nutrients and

growth factors.

Ultrasound Besides mechanical stimulation, ultrasound (US) in the 3-10 MHz range is one of the well-

established therapeutic physical stimuli for bone healing [26]. Ultrasound refers to longitudinal wave

propagation, a special type of sonic wave with a frequency greater than 20 kHz (the upper limit of human

audibility), that causes localized oscillation of particles. Since ultrasound was first reported to stimulate

bone healing in 1950, numerous efforts have been put over several decades to prove its therapeutic

effects in humans [26]–[28]. In particular, low-intensity pulsed ultrasound stimulation (LIPUS), using

intensities less than 50 mW/cm2, has been reported to improve ECM synthesis, accelerate bone healing,

and reactivate failed healing processes [28], [29]. This use of ultrasound to improve bone regeneration

has been approved by the United States Food and Drug Administration (FDA) for human applications since

1994 [30]. In in vitro cell studies, LIPUS was found to enhance the expression of osteoblast maturation

markers, such as osteocalcin (OCN) gene, bone sialoprotein (BSP), and calcium (Ca2+) [31]–[35].

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LIPUS-induced bone healing can occur by the processes of inflammation, soft callus formation,

angiogenesis (development of new blood vessels), early osteogenesis, bone formation, and bone

remodeling [36]. There are several theories to illustrate these mechanisms. In the first theory, the

oscillatory displacement of the cell membrane caused by the ultrasound wave triggers oscillatory

displacement between intracellular elements of different densities [37]. The very low strains induced by

the ultrasound on cells in vitro induces fluidization of the cytoskeleton and acceleration of cytoskeletal

remodeling [38]. Another theory is the bilayer sonophore model, in which the ultrasound application pulls

the cell membrane lipid layers apart and pushes them back together cyclically, leading to

intramembranous hydrophobic spaces expanding and contracting accordingly [39]. In the fourth theory,

ultrasound induces intracellular stress and strain maximized within the cell at two distinct resonant

frequencies. Stimulated load-inducible gene expression, therefore, is maximized when the excitation

frequency matches the cell’s resonant frequency [40]. A final popular theory posits ultrasound can

modulate the microenvironment by heating it, thus regulating thermo-sensitive enzymes like

metalloproteinase, which are important for bone matrix remodeling [41].

Shock Waves Shock waves, a kind of short-duration, acoustic pressure wave with an intensity between 30 MPa

and 100 MPa have also been shown to be an effective form of physical stimulation [42]. These waves can

be produced by various generators including hydraulic, electromagnetic, and pneumatic [42]. After

propagating into the tissue, shock waves result in cavitation, the formation of micro-bubbles in areas of

low pressure on the focal area. Shock waves have been introduced to increase cell membrane

permeability and facilitate the delivery of macromolecules into cells [43]. Clinical use of shock waves,

referred to as extracorporeal shock wave therapy (ESWT), is noninvasive and focused solely on the

treatment area. In bone, ESWT is known to relieve pain, reduce inflammation, induce neo-angiogenesis,

and stimulate stem cell activities thus improving tissue regeneration and healing [44]–[46]. It has been

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shown to restore the healing process in cases of non-unions (a serious complication of bone fractures)

[47], loosen the bone cement during revision arthroplasty (a follow-up procedure to a total knee

replacement) [48], and enhance bone callus formation during bone lengthening [49]. In studies regarding

osteoarthritis (OA), compared to a control group, the ESWT-treated group showed an increased osteocyte

count, higher percentage of subchondral trabecular bone, and greater expression of dentin matrix protein

(DMP1), an osteocyte marker [50]. Increased proliferation and migratory capacity were also shown in

human BMDSCs when exposed to shock waves [51]. ADSCs exposed to ESWT have also shown enhanced

production of osteogenic markers such as RUNX2, alkaline phosphatase (ALP), and mineralized Ca2+

matrix. The mechanisms of shock waves’ effects on bone healing have been found to be related to the

micro-fractures and cavitation they induce [52]. The micro-fractures and cavitation trigger the initiation

of remodeling cycles and neovascularization [46], [53]. Thus, they regulate the growth and maturation of

osteoprogenitor cells, membrane polarization, expression of bone morphogenetic proteins (BMPs), and

activation of the mechano-transduction pathways that are related to acoustic stimulations [44], [54]–[56].

During the mechano-transduction process, mechanosensory components in cell membranes such as

integrins, ion channels, and various sensory and growth factor receptors are activated by shock wave-

induced forces.

Laser Low-level laser therapy (LLLT) with proper doses and output powers has also been reported to

stimulate cellular metabolism, increase protein synthesis, and subsequently enhance bone regeneration

[57]. LLLT has been proven to elevate the structural stiffness of bone callus and increase mitochondrial

respiration and adenosine triphosphate (ATP) synthesis [58]. However, some researchers have also found

LLLT to accelerate bone formation through increased osteoblastic activity, vascularization, and better

organization of collagen fibers [59].

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Introduction to Use of Electrical Physical Stimulation One of the most researched methods of physical stimulation proven effective for bone

regeneration is electrical stimulation (ES). Electrical stimulation treatment is especially fitting given bone

tissue’s unique relationship with electricity. Bone tissue, specifically the collagen-hydroxyapatite matrix

comprising it, has a unique property called piezoelectricity, which allows the tissue to generate electricity

in response to mechanical stress and vice versa [60]. The polar, uniaxial orientation of collagen enables it

to displace its charge carriers from the inside of the molecule to the outside when stressed in an

appropriate way [61]. The exact amplitude of the electrical potential generated, however, is determined

by the rate, magnitude, and orientation of the applied load and the resulting bone deformation. Normally,

when bone is bent, the concave sides (under compressive stress) become negatively charged and the

convex sides (under tensile stress) become positively charged, which makes the bone grow more on the

compressive side and degrade more on the stretched side [62]. Through this same mechanism, stimulation

of bone regeneration is also possible through electrical-induced pathways.

The exact mechanism underlying the intracellular signal transduction of ES in bone repair is still

unclear. Several hypotheses have been reported. (1) ES could alter the ion flux via cell membrane proteins

(such as ion channels, transporters, pumps, and enzymes) and subsequently lead to an ion concentration

change, which may cause depolarization of excitable cells and trigger downstream cellular signaling [63].

(2) Applied current could change the cell gap junctions, which affect the exchange of certain signaling

molecules such as calcium, cyclic nucleotides, and inositol phosphates [64]. Much evidence indicates that

gap junction communication is necessary for the development and maintenance of a differentiated

osteoblast phenotype, including the production of ALP, OCN, bone sialoprotein, and collagen type I (COL

I) [65]. (3) ES may also affect ligand-receptor binding by changing the conformation or expression of

receptors [66]. (4) ES may also stimulate higher metabolic activity, which could induce intracellular ATP

depletion and thus alter the membrane characteristics such as endo- and exocytosis, adhesion, and

Patel 15

motility [67]. (5) ES could change ECM compositions by affecting the ECM components including soluble

ions and charged groups in glycosaminoglycans (GAGs) and proteins [68].

There are three main administrations of electric stimulation. These are summarized in Figure 2 [69].

Two methods produce electrical fields and one method applies a direct electric charge. (a)

Capacitive coupling (CC): Two capacitive coupled electrodes are situated on the skin on both sides of the

treatment site. An external power source is then attached to the electrodes, which induces an electric

field (EF) at the treatment site. (b) Inductive coupling (IC): An electromagnetic current-carrying coil

attached to an external power source is placed on the skin overlying the treatment site. The coil generates

pulsed electromagnetic fields (PEMFs), which induce an electrical field at the site. (c) Direct current (DC):

A cathode is implanted at the treatment site which is attached to either a subcutaneous power source or

an external power source to generate an electric charge at the fracture site. [69]

Capacitive Coupling Capacitive coupling stimulation involves two capacitive coupled electrodes utilizing external

power to induce a physiological electric field around a treatment site. Physiological electric fields serve as

Figure 2. Main Methods of Administering Electric Stimulation to Bone: (a) Capacitive coupling (CC): coupled electrodes situated externally creating an electric field. (b) Inductive coupling (IC): an electromagnetic current-carrying coil situation on the skin generates pulsed electromagnetic fields (PEMFs), which induce an electrical field. (c) Direct current (DC): a cathode is implanted at the treatment site which is attached to either a subcutaneous or an external power source to generate an electric surface charge.

Patel 16

an effective tool to control and adjust cellular and tissue homeostasis. The human body generates a

biological EF ranging between 10 mV and 60 mV at various locations [70]. Bioelectricity is very important

in the wound healing process. When a wound is created, a steady EF is initiated. This endogenous EF

guides cell migration toward the wound edge. In 1953, researchers confirmed this an in in vivo rabbit

model when they applied a continuous electrical current to a rabbit femur for 3 weeks and demonstrated

new bone formation around the cathode [71]. Since then, the use of EFs for bone healing applications has

been widely researched [72].

Inductive Coupling Inductive coupling, utilizing pulsed electromagnetic fields to induce an electrical field at a

treatment site, has added benefits beyond those typically seen with CC [73]. Under IC stimulation,

osteoblasts have been found to exhibit increased osteogenesis caused by elevated expression of

transforming growth factor-beta 1 (TGF-β1) [74] and BMP2/4 [75] and reinforced intracellular Ca2+

transients [76]. In a rat model, long-term IC stimulation treatment alleviated lumbar vertebral

osteoporosis by increasing bone formation and suppressing bone resorption [77]. Researchers have

identified a specific extremely low-frequency pulsed electromagnetic field (ELF-PEMF) in the 10-90.6 Hz

frequency that supports human osteoblast function [78], [79]. The ELF-PEMF increases production of non-

toxic amounts of reactive oxygen species induced anti-oxidative defense mechanisms in these cells [78],

[79]. In bone tissue, ELF-PEMF treatment was found to modulate the cell cycle of MSCs of different origins

to enhance their differentiation and proliferation [80]. This could be seen by their enhanced production

of ECM and growth and differentiation factors including TGF-β1 and BMPs [80]. Besides frequencies that

support osteoblast function, a wide range of electromagnetic stimulation frequencies between 2 Hz and

123 Hz have been shown to be effective in improving osteogenic differentiation in ADSCs [81]. The

stimulation has been found to increase intracellular Ca2+, ALP activity, and cytoskeleton tension after 14

days induction and increase expression of ALP, OPN, RUNX2, and COL I after 21 days induction [81], [82].

Patel 17

Direct Current Direct current has also been observed to enhance osteogenesis when cells are stimulated with a

current of 5–100 μA [83]. Electrical potentials have been proven to play an important role in bone cell

proliferation, migration, and remodeling both in vitro and in vivo [84]–[86]. Some implant materials, such

as electrically active ceramics, including polarized hydroxyapatite and piezoelectric ceramics, have been

found to induce bone growth and improve bone formation around implants respectively [87]. The

mechanism by which direct current influences biological responses is likely to result from preferential

adsorption of proteins and ions onto the charged surface [88].

Numerous studies have emphasized the importance of these surface charges on cell behavior at

the biomaterial interface [84], [85], [89]. In the calvarial bones of rats, after implanting electrically

polarized hydroxyapatite plates, improved bone ingrowth and enhanced osteoblast activity have been

observed, with complete bone penetration into the implants occurring as early as three weeks post-

stimulation application [84]. In another study, bone formation increases occurred on the negatively

charged surfaces of the polarized implants due to a proven accumulation of Ca2+ ions on the negative

surface and adherence of fibronectin, osteocalcin, and BMPs on the positively charged surfaces, thus

improving osteoblast migration [84]. In addition, hyaluronan, an ECM component, has been found to plays

a key role in the cellular interactions between charged surfaces and mediating initial contact between cell

and attachment surfaces [90]. Although much is known about DC use, implementation of surface charge

direct current stimulation treatment has been slow growing.

Introduction to Direct Current Surface Charge Electrical Stimulator Systems Numerous works have shown direct electricity’s ability to promote osteogenesis in ADSC and

BMDSC lines when they are seeded in bone scaffolds [91], [92]. However, typical DC electrical stimulators

are either (a) externally housed and require overcomplex percutaneous wires be connected to the

Patel 18

implanted scaffold or (b) implanted non-degradable devices that contain toxic batteries and require

invasive removal surgeries. Thus, a need exists for new strategies to deliver such stimulation.

Introduction to Use of Piezoelectrics for Surface Charge Electrical Stimulators Piezoelectric materials, a type of smart material that can generate electrical surface charge from

mechanical vibrations such as ultrasound, show much promise in this regard. These materials have been

exploited extensively in industry for important external devices such as sensors, transducers, and

actuators. Piezoelectric materials, however, also offer many significant medical applications when used

inside the body[93]–[95]. For example, piezoelectric pressure transducers have been embedded inside

percutaneous medical catheters to monitor dangerous elevations in physiological pressures including

intracranial pressure[96], blood pressures[97], and bladder pressure[98]. Researchers have also reported

an implantable piezoelectric ultrasonic transducer, which is more advantageous than external

transducers, to disrupt the blood-brain barrier and facilitate the delivery of drugs into the brain [99]–

[101]. With these applications proving its efficacy to transform vibrational energy into electricity,

piezoelectric technology is primed for use as a novel surface charge electrical stimulator for bone

regeneration.

Unfortunately, most commonly-used piezoelectric materials such as lead zirconate titanate (PZT),

polyvinylidene fluoride (PVDF), zinc oxide (ZnO), and barium titanate (BaTiO3), etc. are either toxic and/or

not biodegradable [102]. Implantable devices that utilize these materials require an invasive removal

surgery which can easily damage interfaced tissues and lead to complications, raising a significant safety

concern. The ability to create biodegradable and biocompatible piezoelectric materials can, therefore,

bring about significant medical applications including as an electrical stimulator for bone regeneration.

Introduction to Use of Poly-L-Lactic Acid for Surface Charge Electrical Stimulators Recently, poly-L-lactic acid (PLLA), a biocompatible and biodegradable medical polymer (used for

many FDA-approved implantable devices such as surgical sutures[103], tissue scaffolds[104], and drug-

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delivery carriers[105]), has been shown to exhibit piezoelectricity when appropriately processed, thereby

offering a much safer alternative piezoelectric material [106].

Classical processing of PLLA, which entails stretching an amorphous sheet of the material, and

then annealing it has only yielded mild piezoelectric responses [106]. However, a few researchers have

utilized electrospinning, a nanofiber production method that uses electric force to draw charged threads

of polymer solutions onto a rotating collection plate, to create flexible PLLA nanofiber films which exhibit

a higher level of piezoelectricity [107]–[109]. Typically, piezoelectric PLLA nanofibers struggle with

stability, piezoelectric performance, and reliability [107]–[109]. However recent research in polymer

processing has for the first time allowed the creation of biodegradable and biocompatible piezoelectric

PLLA nanofibers with a highly-controllable, efficient, and stable piezoelectric performance. The PLLA

nanofibers also offer a natural extracellular-matrix like environment to facilitate cellular growth for tissue

regeneration.

Thus here, a new tissue-stimulation approach, using a biodegradable piezoelectric PLLA nanofiber

scaffold in combination with non-invasive ultrasound to generate electrical charges is demonstrated to

enhance osteogenic differentiation of stem cells for eliciting bone-tissue regeneration. This strategy is

analogous to the development of a wireless and battery-free, biodegradable, electrical-stimulator for

bone growth.

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Materials & Methods

Step 1. Processing and Manufacturing of the Piezoelectric PLLA Nanofiber Mesh Unprocessed, PLLA (thought a polar molecule that has inherent charge separation) does not

present with any piezoelectric properties, producing no electricity when mechanically stimulated. In order

to improve the piezoelectric response of PLLA, two major material properties need to be improved to

compound the charge separation to a point that the charge can be released - the crystallinity and the

orientation of the polymer chain. By improving these properties, the carbon-oxygen double bonds present

in the PLLA backbone become aligned resulting in an inherent net polarization and a well-documented

piezoelectric response under an applied force. [110], [111]

The PLLA nanofibers were fabricated using an in-house electrospinning setup shown in Figure 3

[112]. Poly-L-Lactic acid granules (PURASORB PL38, Corbin Purac, Amsterdam, Netherlands) were used in

this work. To create each PLLA mesh, 0.8g of PLLA was dissolved in a 1:4 V/V mixture of N, N –

Dimethylformamide (DMF), anhydrous, ≥99.9%, MilliporeSigma, Burlington, MA) and dichloromethane

(DCM, MilliporeSigma, Burlington, MA) respectively. The polymer solution was placed in a 10‐mL syringe

which was loaded in a syringe pump (New Era Pump Systems Inc., Farmingdale, NY). The solution flow

Figure 3. Piezoelectric PLLA Nanofiber Mesh Manufacturing Diagram: (a) Electrospinning setup with a 1:4 DMF:DCM v/v PLLA solution loaded into the syringe, a 14kV power source drawing out the solution, a rotating collection drum with an adjustable rotation speed between 300 and 4000 rpm (b) Single, final PLLA nanofiber mesh

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rate was 2 mL/hr through a flat-tipped 22-gauge needle (Jensen Global, Santa Barbara, CA) with 14kV

applied to it. A grounded, aluminum rotating drum was positioned 8.6 cm from the needle tip, and a

voltage potential was applied to the needle using a high voltage DC power supply (Gamma High Voltage,

Ormond Beach, FL). The polarized solution was then sprayed at a grounded aluminum drum, wrapped in

aluminum foil, rotating at speeds from 300 – 4000 rpm (rotations per minute). The relative humidity was

monitored using a NIST‐certified hygrometer (Cole-Parmer, Vernon Hills, IL). The experiments were

conducted in a 40 ±15 % relative humidity atmosphere at ambient temperature.

This resulted in a PLLA nanofiber mat with varying degrees of alignment and fiber diameter, that

varied with rotating drum speed. The nanofiber samples initially made by the electrospinning setup were

expectedly highly amorphous. Therefore, the samples were annealed to further improve the material’s

crystallinity. The samples were initially annealed at 105°C for 10 hours to initiate pre-crystallization and

then slowly cooled to room temperature. Then the samples were annealed between Teflon FEP sheets

(American DURAFILM, Holliston, MA) and glass slides at 160.1°C for 10 hours and again slowly cooled to

room temperature.

After the production of the PLLA nanofiber meshes, visualization of the films occurred via scanning

electron microscopy (SEM). For each sample, a square PLLA film with dimensions 7mm x 7mm was

mounted on a standard SEM pin (Ted Pella, Redding, CA) using carbon conductive tabs (Ted Pella, Redding,

CA). The sample was then coated in gold-palladium for 1 minute using a sputter coater (Polaron E5100,

Quorum Technologies Ltd., Lewes, UK). The sample was finally imaged using an FEI TeneoLoVac SEM at

5kV and 2,500x magnification. These images can be seen in Figure 7 and Figure 8. These studies were

performed at the University of Connecticut/Thermo Fisher Scientific Center for Advanced Microscopy and

Materials Analysis).

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With the films prepared, characterization of the films and verification of their piezoelectric

efficiency (their ability to convert vibrational energy into electrical energy) was tested. The crystallinity of

the PLLA nanofiber samples was analyzed using differential scanning calorimetry (DSC). The orientation

of crystalline domains was visualized via 1-Dimensional & 2-Dimensional X-ray diffraction (XRD).

Step 2. Verification of PLLA Nanofiber Mesh Efficacy and Degradation To verify the ability of the PLLA nanofiber mesh to generate surface charge in aqueous in vitro

conditions, a test was conducted in which the PLLA mesh was submerged in solution, ultrasonically

stimulated, and the electrical output was measured. To set up this experiment, first, gold electrodes were

electrodeposited on two sides of a 1cm x 1cm square-cut PLLA mesh piece. Following this, they were

covered by polydimethylsiloxane (PDMS) to waterproof the metal electrodes and then connected to

PDMS-encapsulated copper wires. Gold electrodes were used to reduce the noise produced by the PLLA

nanofiber. The PLLA with electrodes was then placed inside a medium of phosphate buffer solution (PBS)

inside a sonication bath (Branson CPX 2800, Emerson Electric Co., St. Louis, MO). The PLLA mesh was

under PBS while the two copper wires were connected to an electrometer to output the measured charge

from the PLLA. As the surface charge outputted was dependent on the ultrasound stimulation, a range of

ultrasound frequencies with different intensities was applied on top of the submerged PLLA mesh to

engineer the piezoelectric outputs and optimize regenerative outcomes. Ultimately, the PLLA was

stimulated with 40kHz 0.4W/cm2 ultrasound. The results of this can be seen in Figure 9.

After verifying the piezoelectric capabilities of the processed PLLA nanofiber, a degradation study

was performed to make sure that the electrospun PLLA follows a similar degradation as traditionally

stretched PLLA, which has already been verified by the FDA to follow sufficient degradation standards.

The experiment was performed, following a previous report [98]. The nanofiber meshes (n = 3 for each

PLLA film type) were initially placed in PBS at 37°C. As PLLA has a long degradation time (about 180 days),

a slightly sped up degradation study occurred [113]. From days 1-28, the PBS was maintained at a

Patel 23

physiological temperature of 37°C. From days 28 to 40 however, the PBS was doubled to 74°C. The results

of the degradation study can be found in Figure 10.

Step 3. Assembly of the PLLA Scaffold After verifying the single layer efficiency and degradation of the PLLA nanofiber mesh, a multi-

layered bone scaffold was produced. A diagram of this bone graft can be seen in Figure 4. This graft was

produced by simply layering three Piezoelectric PLLA meshes alternated by layers of stem cells suspended

in cell culture media.

Ultimately, two main scaffolds were produced for the in vitro system test for osteogenic

differentiation. The scaffolds used as the experimental group were spun at 3000 rpm and 1000 rpm. These

scaffolds were compared to scaffolds spun at 300 rpm which served as a negative control group.

Step 4. Test of Osteogenic Differentiation in vitro In the in vitro experiment, ADSCs and later BMDSCs were seeded on experimental piezoelectric

PLLA films (3000 rpm and 1000 rpm films) and control, non-piezoelectric films (300 rpm films). These

seeded scaffolds were then stimulated with 40 kHz, 0.4W/cm2 US for 20 minutes/day for 10 days to study

osteogenesis. Five different experimental groups described in Figure 5 were used to test the efficacy of

the entire system on enhanced osteogenesis.

Figure 4. Diagram of the Completed PLLA Bone Graft: (macro image) The multi-layered scaffold seeded with stem cells. (micro image) A rat model showing a theoretical implantation and ultrasound stimulation site in a rat bone defect.

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Prior to the start of the experiment, sterilization of the PLLA scaffolds and preparation of the cell

culture plates occurred. The 1cm x 1cm scaffolds were sterilized using ethanol and UV treatment. The

entire process was carried out under a Class II laminar flow cell culture hood. First, the scaffolds were

soaked in 70% ethanol for 30 minutes. Then, the ethanol was removed, and the scaffolds were placed

under UV light for 20 minutes on each side. This made the PLLA scaffolds suitable for cell seeding.

Post sterilization, the scaffolds were fixed onto 6-well culture plates (Thermo Scientific) using

biocompatible silicone glue (KWIK-SIL, World Precision Instruments). A small amount of glue was added

to each well, spread across the entire surface using a spatula, and used to adhere the PLLA nanofiber

scaffold. The glue was used to keep the scaffolds securely on the plate while the US treatments were

carried out and used to prevent the ultrasound waves from getting reflected around by the walls of the

polystyrene wells and getting dissipated in the process. After that, the plates were set aside to allow the

glue to dry and the scaffolds to be secured in the plates.

Cells were seeded onto the PLLA scaffolds after the latter were sterilized and glued to the plates.

The cells used for this purpose were adipose-derived stem cells (XCells Biotechnologies) and were

purchased at passage one. The cells, received in frozen, pelleted form, were thawed, plated, and

expanded until passage five, as per the company protocol. For expanding the cultures, proliferation media

Figure 5. Table of Experimental Groups: Group 1) 3000 rpm film treated with ultrasound. Group 2) 1000 rpm film treated with ultrasound. Group 3) 300 rpm film treated with ultrasound. Group 4) 3000 rpm film not treated with ultrasound. Group 5) 300 rpm film not treated with ultrasound.

Patel 25

with 10 % fetal bovine serum and 1 % penicillin-streptomycin antibiotic solution was prepared in

Dulbecco`s Modified Eagle Media (DMEM) (all three components purchased from Gibco). Expansion was

carried out by passaging the cells every 5-6 days after the culture reached 80-90 % confluence. Ultimately,

at 80-90 % confluence of passage five cells, they were detached from the flasks using Trypsin /

Ethylenediaminetetraacetic acid (EDTA) (purchased from Gibco) and seeded onto the PLLA scaffolds at a

seeding density of 5 x 105 cells per scaffold. The cells were allowed to attach for one day under

proliferation media. After that, the 2mL of proliferation medium was replaced with an equal volume of

osteogenic differentiation medium that was prepared by adding 50 µg/ml ascorbic acid and 10 mM beta-

glycerophosphate to the proliferation medium prepared previously [CITATION]. The cultures were left in

osteogenic differentiation medium for one day before we started the daily ultrasonic treatments.

After one day in differentiation media, the cells and scaffold began their 10-day US stimulation

treatment. A diagram of this setup is seen in Figure 6.

The treatment was performed using a sonication cleaning bath (Branson CPX 2800, Emerson

Electric Co., St. Louis, MO). The ultrasound provided was at 40 kHz and 0.4W/cm2. The treatment was

done for 20 minutes each day and performed for 10 days. First, the cell culture plate was sealed in a

Figure 6. Diagram of Ultrasound Stimulation Setup: Schematic of the cell culture plate and ultrasound stimulation setup used for in vitro experiments. US was applied to every cell culture plate at 40kHz and 0.4W/cm2 for 20minutes/day for 10 days. Various piezoelectric PLLA films (3000 rpm and 1000 rpm meshes) and non-piezoelectric films (300 rpm meshes) were seeded with ADSCs or BMDSCs and stimulated with US to test the effect of surface charge on osteogenesis.

Patel 26

standardized manner. First, we taped the lid onto 4 sides of the plate using labeling tape. Then we

removed the plate from the cell hood and encapsulated it in two layers each of plastic wrap (Kirkland

Signature Stretch-Tite Plastic Wrap - 11 7/8 x750 Feet) and duct tape (manufactured by 3M). Three layers

of plastic wrap were first applied in alternating orientations followed by a single layer of duct tape. Lastly,

a centrally-placed tab was created on the top of the encapsulated plate to suspend the plate in the

sonication water bath. This was done by using a laboratory clamp, stand apparatus, and an alligator clip.

The plate was suspended so that it was submerged halfway into the water and horizontally level. The plate

was sonicated for 20 minutes. When the treatment was complete for the day, the plate was removed

from the bath, encapsulation layer removed, cell culture media change, and returned to the incubator

until the next US treatment.

Step 5. Assessment of Cell Proliferation and Differentiation After US treatment, three main cell proliferation and differentiation tests were conducted: an ALP

quantification assay, an alizarin red quantification assay for CA2+ mineralization, and a quantitative

polymerase chain reaction (qPCR) for gene expression measurement. After 10 days of treatment, the

cultures were terminated and analyzed for osteogenic differentiation activity.

ALP is an enzyme released by osteoblasts and pre-osteoblasts; its presence thus indicates the

osteogenic differentiation of ADSCs. ALP activity measurement was carried out using a p-nitrophenyl

phosphate (pNPP) based quantification kit (purchased from Bio-Rad, cat no-172-1063). The pNPP solution

was prepared and added to the cellular protein extract and the ALP activity levels were quantified in

accordance with the kit protocol. The standard curve for ALP-pNPP product quantification was prepared

by dissolving different amounts of ALP (purchased from Sigma-Aldrich) in an aqueous solution, reacting

them to pNPP, and reading the absorbance of each solution. This helped quantify the absolute ALP in our

cultures. The values obtained were normalized by total protein content. The ALP data was normalized

using total protein content measured using the bicinchoninic acid (BCA) assay (purchased from Pierce™).

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The BCA assay (kit purchased from Pierce™, Thermo Scientific) was used to quantify the total

protein content of the cultures. As per the kit protocol, a protein solution was extracted from each culture

and was used to normalize the ALP assay described prior and the mineralization assay described below. A

one-way analysis of variance (ANOVA) test and a Tukey’s Honest Significant Difference (HSD) test was

used to assess significance.

As mineral deposition, especially of Ca2+, is a common property of bone cells, the alizarin red assay

was used to quantify the amount of the Ca2+ mineral formed by the cells seeded in each group. For the

alizarin red assay, the cultures were fixed in 70% ethanol at 4°C for one hour. Following this, the ethanol

was removed, the wells were rinsed and the alizarin red dye (purchased from EMD Millipore corp.) was

added and the cultures were incubated at room temperature for 30 minutes. After this incubation period,

the dye was rinsed away, and the wells were viewed under the microscope to observe the then stained

red mineral formation. Quantification of the red mineral formation was done by de-staining the red dye

in cetyl-pyridinium chloride (purchased from Sigma Aldrich), measuring its absorbance, and equating that

to the absolute concentration of Ca2+ produced. The values obtained were lastly normalized by total

protein content using the BCA assay described prior.

To measure the expression of osteocalcin and osterix genes, which are typical osteogenic

differentiation markers, qPCR was conducted. qPCR quantification was performed using the universal sybr

green master mix (purchased from Bio-Rad). The forward primer used for osterix was: 5’-GGA AAG GAG

GCA CAA AGA A-3’ and the reverse sequence used was 5’-GTC CAT TGG TGC TTG AGA A-3’. The forward

primer used for osteocalcin was 5’-CAA GCA GGA GGG CAA TAA G-3’ and the reverse primer was 5’-CGT

CAC AAG CAG GGT TAA G-3’ (purchased from Integrated DNA Technologies). The corresponding protocol

for the qPCR kit and primers was used to guide the qPCR process. Extraction of RNA was performed using

the Trizol reagent, the extracted RNA was converted to cDNA using the iScript cDNA synthesis kit

(purchased from Bio-rad), the cDNA was mixed with the appropriate primers and the master mix and run

Patel 28

through an RT-qPCR machine (purchased from ABsystems) to get the gene expression values. These values

were normalized with a qPCR assay of beta-actin, a common housekeeping gene. The beta-actin forward

primer used was 5’-TCC TCC TGA GCG CAA GTA CTC T-3’ and the reverse primer was 5’-CGG ACT CAT CGT

ACT CGT GCT T-3’). A one-way ANOVA test and a Tukey’s HSD test was used to assess significance.

In addition to ADSCs, the ability of the piezoelectric nanofiber scaffold under an applied US to

induce osteogenesis from bone marrow stem cells was also investigated. Specifically, a fluorescent

reporter BMDSC system was utilized to monitor the proliferation and differentiation of our cells in real-

time and to confirm the osteogenic properties of our PLLA system. The reporter cells used were primary

BMDSCs with a topaz green fluorescent protein reporter for bone sialoprotein (BSP-GFP-topaz) and a

mCherry red fluorescent protein reporter for dentin matrix protein (DMP1-RFP-mCherry) tag. This cell line

was harvested from the tibial and femoral bone marrows of 3 to 4-week-old dual transgenic mice

containing BSP-GFP-topaz and DMP1-RFP-mCherry fluorescent reporter genes.

The progressive expression from BSP to DMP was used to determine the different stages of

differentiation BMDSCs go through. Optical imaging was used to visually compare the osteogenic

properties of different scaffold types and treatments. At days 0 (pre-seeding), 1, 2, and 3 of US treatment,

reporter cell fluorescence (n=3) was captured using the Zeiss Axio Observer Z.1 inverted fluorescence

microscope. The fluorescence of the cells was captured by taking sectional images of the entire scaffold

surface in ZEN software at an objective of 5X. Exposure times of 100 ms and 1000 ms were applied for

both the BSP-GFP-topaz and the DMP1-RFP-mCherry channels. The sectional images obtained were then

stitched together to create a mosaic. The final images obtained showed different amounts of green and

red fluorescence for different scaffolds receiving different treatments. The amount of fluorescence in each

image was quantified using the ImageJ software and the relative fluorescence of each group was

compared. A one-way ANOVA test and a Tukey ‘s HSD test was used to assess significance.

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Results

Step 1. Processing and Manufacturing of the Piezoelectric PLLA Nanofiber Mesh As the speed of the rotating drum was varied from 300 rpm – 4000 rpm, the PLLA nanofiber films

were produced with different levels of fiber orientation. This can be seen in Figure 7 and Figure 8.

s

s

Figure 7. SEM Images of Non-Piezoelectric PLLA Nanofiber Film: 300 rpm Non-Piezoelectric PLLA film confirming the poor orientation of the nanofibers.

Figure 8. SEM Images of Piezoelectric PLLA Nanofibers: PLLA films produced at 1000 rpm – 4000 rpm, confirming higher electrospinning rotation speed increases fiber alignment. Scale bars are all 40 µm.

Patel 30

Step 2 & 3. Verification of PLLA Nanofiber Mesh Efficacy and Degradation & Assembly of

the PLLA Scaffold Ultimately, the 3000 rpm film was proven to output +/- 40mV of electrical surface charge when

stimulated with 40kHz 0.4W/cm2 ultrasound. The untreated, 300 rpm film shows only noise and has no

change in electrical output when US is applied. This can be seen in Figure 9.

With the piezoelectric efficacy of the experimental 3000 rpm film confirmed, a degradation study

was done on the PLLA nanofiber. The graft of this degradation can be seen in Figure 10.

Figure 10. Graph of PLLA Degradation Study: Remaining weight (%) of piezoelectric PLLA films degraded in PBS for 40 days. Initially, the temperature of the PBS was a physiologic 37°C. On day 28, this was doubled to 74°C to speed up degradation.

Figure 9. Verification of PLLA Piezoelectric Output: (Right) Schematic of PLLA scaffold testing done in a salt bath with electrical output read by an oscilloscope. (Left) Piezoelectric voltage output when 40kHz ultrasound is off versus on.

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Step 4 & 5. Test of Osteogenic Differentiation in vitro & Assessment of Cell Proliferation

and Differentiation On day 10 of the cell culture tests, three assays were done on the ADSCs and the BMDSCs. The

results of these assays are below. In total, there were 5 groups, which can be seen in Figure 5. The results

of the ADSCs assays can be seen in Figure 11 and Figure 12.

The results of the subsequent gene expression assays for the ADSCs are displayed in Figure 12.

Following ADSC studies, reporter BMDSC studies were conducted. Figure 13 shows the results of

the assessment of BMDSC reporter differentiation. Figure 13a shows the differentiation pathway from

progenitor stem cells to differentiated osteocyte. Figure 13b consists of fluorescent images of the cells in

different experimental conditions. Figure 13c shows the florescent expressions quantitively and provides

a graphical comparison between each group.

Figure 11. ADSC Protein and Mineral Expression Test Results: (a) Quantitative comparison of ALP production by the cells in different treatment groups. (b). Comparison of Ca2+ production by the cells in different treatment groups. A one-way ANOVA and a Tukey’s HSD test was used as a test of significance. In the figure, † represents a significance level of 0.01, †† represents a significance level of 0.001, and ††† represents a significance level of 0.0001.

Figure 12. ADSC Gene Expression Test Results: (a) Osteocalcin gene levels by the cells in different treatment groups (b) Osterix gene levels by the cells in different treatment groups. Both genes levels were normalized to the gene expression of beta-actin. A one-way ANOVA and a Tukey’s HSD test was used as a test of significance. In the figure, † represents a significance level of 0.01, †† represents a significance level of 0.001, and ††† represents a significance level of 0.0001.

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Figure 13. BMDSC Reporter Cell Activity: This figure shows the florescent expressions of primary bone marrow stem cells (BMDSCs) with reporters for bone sialoprotein (BSP-GFP-topaz) and dentin matrix protein (DMP1-RFP-mCherry) when seeded on electrospun PLLA scaffolds and exposed to US treatment. These reporter stem-cells fluoresce green and red when differentiated into osteoblasts and osteocytes respectively, two signs of eliciting bone regeneration. (a) Schematic of the progressive expression from Stem Cell to differentiated Osteocyte. (b) Fluorescent microscopy of the cells seeded on different scaffolds in different treatment groups through the GFP channel (100ms exposure time) and the cherry red channel (1000ms exposure time). (c) Quantitative fluorescent expressions and graphical comparison of between each group. A one-way ANOVA and a Tukey’s HSD test was used as a test of significance. In the figure, † represents a significance level of 0.01, †† represents a significance level of 0.001, and ††† represents a significance level of 0.0001.

Patel 33

Discussion

Ultimately, as expected, nanofibers from were made with varying piezoelectricity, based on the

rotational speed of the electrospinning collecting drum. As can be seen in Figure 7 and Figure 8, the

nanofiber films, collected at slow spin-speeds, have lower levels of fiber alignment, thus offering lower

piezoelectric effects. Past literature has reported a positive relationship between electrospinning

collecting drum speed and crystallinity of the nanofiber material [104]. Consistently, as seen in Figure 8,

the 3000 rpm and 4000 rpm samples have the best fiber alignment. Higher speeds were not tested due

to reported strain overload on the fibers that can break them when spun at such high speeds [104]. For

this reason, the 3000 rpm film was chosen as the positive, experimental PLLA control group and the 300

rpm film was used for the negative control group.

When the 3000 rpm film was tested in a salt-bath solution, it was proven to output +/- 40mV of

electrical surface charge when stimulated with 40kHz 0.4W/cm2 ultrasound. This can be seen in Figure 9.

Also in Figure 9, however, is significant noise from the non-piezoelectric 300 rpm film. Earlier studies of

PLLA electrospun films produced at this speed have shown that they are not oriented enough to allow for

sufficient charge polarization to create usable electricity, still significant noise was seen [109]. Related

research on PLLA stimulation has found that PLLA also exhibits another electricity-making effect called the

triboelectric effect [114]–[116]. Triboelectric materials become electrically charged after they are

separated from a previously-in-contact different material [114]–[116]. The filtered, useful +/- 40mV

voltage falls within a clinically proven range of electrical stimulation that has been proven to be beneficial

for bone regeneration [66].

With the piezoelectric efficacy of the experimental 3000 rpm film confirmed, a degradation study

was done on the PLLA nanofiber to confirm its degradation matched that of the FDA approved, stretched

PLLA polymer. The graft of this degradation can be seen in Figure 10. Although standard PLLA has a proven

Patel 34

degradation time of approximately 180 days, a 40-day study was conducted here. Over the course of the

first 28 days, there was minimal change in the weight of all films [117]. To illustrate the influence of

processing condition on degradation, the temperature was increased to 74˚C on day 28 to accelerate the

film erosion. Still, both the standard, FDA-approved stretched compression-molded PLLA film and the

electrospun sample, collected at 3000 rpm, degrade at similar rates. By day 40, both samples were at

approximately 60% of their original weight. While this is the result of an accelerated degradation, the

trend is expected to hold for degradation studies performed over a longer period of time at 37˚C. Optimal

degradation studies have a constant downward trend, reflective of the fact that once degraded, a material

cannot be rebuild. Figure 10 data however has an upward spike on day 20 and day 32, representing a

theoretical gain in mass of the PLLA nanofiber. While water absorption into a sample has been shown to

result in this [118], it should not be occurring in a material such as PLLA. Again, this provides another point

of investigation.

Following verification of efficiency and degradation for the PLLA, the entire system was tested.

Figure 5 shows all the experimental groups utilized. Figure 11 shows the protein and mineralization

expression of these groups and Figure 12 shows the gene expression of these groups. In Figure 11a, the

groups of 3000 rpm PLLA + US and 1000 rpm PLLA + US have significantly higher (p<0.0001) expressions

of ALP than the other three groups. In addition, the 3000 rpm + US (with more piezoelectric charge) has

significantly higher ALP production compared to the 1000 rpm + US (with less piezoelectric charge)

(p<0.001). Figure 11b demonstrates that the groups of 3000 rpm PLLA + US and 1000 rpm PLLA + US have

significantly higher (p<0.0001) mineral formation as compared to the other sham/control groups. Similar

to the ALP assay, the 3000 rpm + US group has significantly higher mineral formation when compared to

the 1000 rpm + US group (p<0.001). Figure 12a and Figure 12b show that the 3000 rpm and 1000 rpm

PLLA + US show significantly higher levels of both osteocalcin (p<0.0001) and osterix (p<0.0001)

expression respectively when compared to the other groups. Again, the 3000 rpm PLLA + US group has

Patel 35

significantly higher expression of both genes as compared to the 1000 rpm + US group, illustrating that

more surface charge generates more osteogenesis. These findings support current findings that mild

direct current electrical stimulation increases stem cell ALP production and Ca2+ production [66].

Reporter BMDSC studies were conducted to measure the active expression of proteins in culture.

Figure 13b shows different experimental groups receiving different treatments as observed under the BSP

mCherry channel and DMP1 cherry red channel. The increase in green and red fluorescence progressively

from the non-piezoelectric 300 rpm PLLA films to the highly piezoelectric 3000 rpm PLLA films shows the

clear increase in osteogenesis as surface charge increases and approaches +/- 40mV. In Figure 13c, it can

be seen that the BMDSC reporter genes are turned on significantly when the cells are cultured on the

piezoelectric 1000 rpm and 3000 rpm PLLA nanofiber scaffolds under applied US, compared to the other

control/sham samples. This result, like the ADSCs presented in Figure 11, demonstrates the effect of

piezoelectric charge on osteogenic differentiation of stem cells.

The fact that PLLA was successfully used in combination with externally applied ultrasound to

produce an electric surface charge that could stimulate stem cell osteogenesis is extremely significant to

the field of bone regeneration. Prior to this novel surface charge electrical stimulation setup, electrical

stimulators relied on externally housed power sources or implanted non-biodegradable batteries [69].

The prior, typically required overcomplex and uncomfortable percutaneous wires to be connected to the

implanted scaffold and the latter, the battery, often contained toxic chemicals and required invasive

removal surgeries. With a safe method of surface charge, electrical stimulation now proved, one hurdle

slowing the implementation of surface charge stimulation for treatment of bone fractures and bone

defects has been overcome [69].

Still, there is much that can be improved about this PLLA nanofiber – ultrasound system. Current

manufacturing methods for electrospun PLLA limit the level of alignment and thus the level of

Patel 36

piezoelectricity that can be achieved [104]. While this processing method is certainly superior to the

classic drawing and annealing method, it still does not yield a piezoelectric response as significant as some

of the classic piezoelectric materials (PZT, PVDF, ZnO, or BaTiO3) [102]. Improved future processing of PLLA

could allow for biodegradable implants that output ever greater levels of surface charge – levels that may

be clinically significant to other cell lines and for use in other treatments [119].

Besides the piezoelectric processing method of PLLA, the 3D scaffold structure is another area for

future investigation. Current technologies limit us to simple stacking of piezoelectric PLLA, however,

research has found that more complex 3D structures can elicit even greater stem cell proliferation and

differentiation [120]. This provides another avenue for the improvement of this PLLA-ultrasound system.

On the ultrasound end, in this study, stimulation only occurred from a single, inferior direction.

However, stimulating a 3D scaffold requires multidimensional ultrasound stimulation [121]. This again

offers another opportunity to increase the achieved piezoelectric output from a PLLA scaffold.

Despite all these potential future directions, there is unknown about the long-term benefits or

risks of ultrasound stimulation and electrical stimulation [122]. As in vivo research has not yet been

conducted, tissue damage from extended ultrasound application or from escaping electrical charge has

not been studied. Additionally, the effect of said ultrasound and resulting electricity may be a hindrance

to vascular flow and miscellaneous cellular growth [6]. These limitations will have to be investigated in

follow up studies.

Overall, this advancement helps make tissue engineering a treatment option that can be ever more

broadly applied. The development of a biodegradable electric stimulator has many applications outside

of bone growth generators. This system could be used to power and wirelessly charge implantable

batteries, such that they never need to be invasively extracted or replaced. Neurostimulators that use

small electrical shocks to disrupting certain signals (often pain signals) traveling through the brain could

Patel 37

be one potential benefactor of this technology [123]. Pacemakers and implantable cardiac defibrillators

are another [123]. Implementation of a piezoelectric PLLA scaffold could be used to harness external

ultrasound vibrations and output them as electrical current to power these medical implants. Without the

need for these invasive battery replacement surgeries or the need for percutaneous leads to be inserted,

fewer complications from these surgeries will occur, fewer resources are needed as no follow-up surgery

is required, and most important patient lives will be benefited.

The novel biodegradable PLLA piezoelectric device described herein is expected to have a profound

impact not only in bone regeneration but in the entire fields of medical implants and tissue engineering.

Patel 38

Conclusion

Ultimately, the production of surface-level electrical charge from ultrasonic stimulation of PLLA

and the use of that charge to promote osteogenic differentiation in ADSC and reporter BMDSC lines was

successfully demonstrated. The assays of both ADSCs and reporter BMDSCs demonstrate this

achievement from a cellular activity standpoint, with both of these differentiated cell lines successfully

carrying out their typical roles in bone growth and repair.

This research presents a novel biodegradable piezoelectric PLLA nanofiber scaffold with stable,

effective, and highly controllable piezoelectric performance that uses only common biodegradable and

biocompatible materials. The PLLA nanofibers and the novel biodegradable piezoelectric device described

herein are expected to have a profound impact on the use of piezoelectrics for surface charge stimulation

in bone regeneration.

Patel 39

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