Development of l-Amino-Acid-Based Hydroxyl Functionalized Biodegradable Amphiphilic Polyesters and Their Drug Delivery Capabilities to Cancer Cells

Article abstract of DOI:10.1021/acs.biomac.9b01124

Hydroxyl-functionalized amphiphilic polyesters based on l-amino acid bioresources were designed and developed, and their nanoassemblies were explored as intracellular enzyme-biodegradable scaffolds for delivering anticancer drugs and fluorophores to cancer cells. To accomplish this task, acetal-masked multifunctional dicarboxylic ester monomer from l-aspartic acid was tailor-made, and it was subjected to solvent-free melt transesterification polycondensation with commercial diols to produce acetal-functionalized polyesters. Acid-catalyzed postpolymerization deprotection of these acetal-polyesters produced amphiphilic hydroxyl-functionalized polyesters. The amphiphilic polyesters were self-assembled in aqueous medium to produce nanoparticles of size <200 nm. Wide ranges of both water-soluble and water-insoluble anticancer drugs such as doxorubicin (DOX), camptothecin (CPT), and curcumin (CUR) and fluorophores such as Nile red (NR), Rose Bengal (RB), and Congo red (CR) were encapsulated in hydroxyl polyesters nanoparticles. In vitro drug release studies revealed that the aliphatic polyester backbone underwent lysosomal enzymatic-biodegradation to release the loaded cargoes at the intracellular compartments. Lysotracker-assisted live-cell confocal microscopy studies further confirmed the colocalization of the polymer nanoscaffolds in the lysosomes and supported their enzymatic-biodegradation for drug delivery. In vitro cytotoxicity studies showed that the nascent polymers were not toxic, whereas their anticancer drug-loaded nanoparticles exhibited excellent cell killing in cervical cancer (HeLa) cell lines. The drug-loaded (CPT, CUR, and DOX) and the fluorophore-loaded (NR, RB, and CR) polymer nanoparticles were highly luminescent; thus, the encapsulated polymer nanoparticles enabled the multiple color-tunable bioimaging in cancer cells in the entire visible region from blue to deep red. Time-dependent live-cell confocal microscopy studies established that the cellular uptake of drugs and fluorophores was 5 to 10-fold higher while they were delivered from the hydroxyl polyester platform. The hydroxyl polyester nanocarrier design strategy opens up new opportunities in drug delivery to cancer cells from a biodegradable polymer platform based on l-amino acids.

Full text of DOI:10.1021/acs.biomac.9b01124

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Article
Development of L-Amino Acid Based Hydroxyl
Functionalized Biodegradable Amphiphilic Polyesters
and Their Drug Delivery Capabilities to Cancer Cells
Sonashree Saxena, and Manickam Jayakannan
Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.9b01124 • Publication Date (Web): 08 Oct 2019
Downloaded from pubs.acs.org on October 13, 2019
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Development of L-Amino Acid Based Hydroxyl
Functionalized Biodegradable Amphiphilic
Polyesters and Their Drug Delivery Capabilities to
Cancer Cells
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Sonashree Saxena and Manickam Jayakannan*
Department of Chemistry
Indian Institute of Science Education and Research (IISER), Pune
Dr. Homi Bhabha Road
Pune 411008, Maharashtra, India
Corresponding Author: E-mail: jayakannan@iiserpune.ac.in
Keywords
Functional Polyesters, L-Amino acids, Enzyme-responsive, Biodegradable polyesters, Nano-
assemblies, Drug Delivery.
Abstract
Hydroxyl-functionalized amphiphilic polyesters based on L-amino acid bio-resources were
designed and developed and their nano-assemblies were explored as intracellular enzyme-
biodegradable scaffolds for delivering anticancer drugs and fluorophores to cancer cells. To
accomplish this task, acetal-masked multi-functional dicarboxylic ester monomer from L-
aspartic acid was tailor-made and it was subjected to solvent-free melt transesterification
polycondensation with commercial diols to produce acetal-functionalized polyesters. Acid-
catalyzed post-polymerization de-protection of these acetal-polyesters produced amphiphilic
hydroxyl-functionalized polyesters. The amphiphilic polyesters were self-assembled in
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aqueous medium to produce nanoparticles of size < 200 nm. Wide ranges of both water-
soluble and water-insoluble anticancer drugs such as doxorubicin (DOX), camptothecin
(CPT) and curcumin (CUR) and fluorophores such as Nile red (NR), Rose Bengal (RB) and
Congo red (CR) were encapsulated in hydroxyl polyesters nanoparticles. In vitro drug release
studies revealed that the aliphatic polyester backbone underwent lysosomal enzymatic-
biodegradation to release the loaded cargoes at the intracellular compartments. Lysotracker
assisted live-cell confocal microscopy studies further confirmed the co-localization of the
polymer nano-scaffolds in the lysosomes and supported their enzymatic-biodegradation for
drug delivery. In vitro cytotoxicity studies showed that the nascent polymers were not toxic
whereas their anticancer drug-loaded nanoparticles exhibited excellent cell killing in cervical
cancer (HeLa) cell lines. The drug (CPT, CUR, and DOX) and the fluorophores (NR, RB,
and CR) loaded polymer nanoparticles were highly luminescent; thus, the encapsulated
polymer nanoparticles enabled the multiple color-tunable bio-imaging in cancer cells in the
entire visible region from blue to deep red. Time-dependent live-cell confocal microscopy
studies established that the cellular uptake of drugs and fluorophores were accomplished 5 to
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0 folds higher while they were delivered from the hydroxyl polyester platform. The
hydroxyl polyester nano-carrier design strategy opens up new opportunities in drug delivery
to cancer cells from biodegradable polymer platform based on L-amino acids.
Introduction
Synthetic polymers based on bio-resources are emerging as very important biomaterials due
to their excellent biocompatibility under physiological conditions and the availability of the
abundant raw materials, and so on so forth.1-4 L-Amino acid bio-resources have unique
features in terms of functional group diversity such as carboxylic acid, amine, hydroxyl, aryl
substitution, phenolic, heterocyclic, etc; thus, they provide unlimited opportunities to make
bio-degradable and biocompatible polymeric materials.5-8 Ring-opening polymerization
(ROP) is one of the widely employed synthetic routes for L-amino acid resources to make
high molecular weight neutral and charged polypeptides,9-11 di- and tri-block copolymers,12-13
poly(-hydroxy acid)s,14-15 etc. These synthetic polypeptides exhibited both -helix and -
sheet secondary structures as similar to that of proteins. The polypeptides were explored for
biomedical applications in drug and gene delivery, tissue engineering, and reinforced nano-
composites in bone replacement, etc.16-18 In the last two decades polycondensation
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approaches have been developed to construct non-peptide polymer analogs from L-amino
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acid resources for biomedical application. Some of the most important non-peptide polymers
are poly(ester-amide)s,19-23 polycarbonates,24-29 poly(ester-urethane-urea),30-33 poly(ester-
urea),34-39 poly(disulfide-urethane)s,40-42 poly(acetal-urethane) etc. We have reported melt
polycondensation route to make linear poly(ester-urethane)s44-46 hyperbranched poly(ester-
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48-49
urethane)s, amine and disulfide functionalized polyesters,
pH and enzyme-responsive
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aliphatic polyesters and thermo-responsive poly(ester-urethane)s and polyurethanes, etc.
Lysosomal enzymes like esterase, trypsin, and -chymotrypsin, etc were found to be efficient
in the biodegradation of the non-peptide polymer nano-assemblies under physiological
conditions.53,54 Enzyme-responsiveness was explored as a trigger in delivery of anticancer
drugs,23,29,40,50-52 FRET-based bio-imaging,54-55 and tissue-fillers,56-58, etc. Unambiguously, the
amphiphilicity in both peptide and non-peptide polymer systems were largely accomplished
by the introduction of polyethylene glycols (PEG) in the polymer matrix. Though PEG-unit is
an excellent choice in nano- design; they are non-functional which restrict their feasibility for
anchoring targeted ligands and drugs, etc until otherwise tedious and multi-step synthetic
efforts are taken. This emphasized that the design and development of functional
biodegradable polymers from L-amino acid resources with appropriate solubility (or
dispersibility) in aqueous medium could open new generation of polymer structures for their
long-term utility in the biomedical field. Unfortunately, functional non-peptide polymers
from L-amino acid is one of the least explored areas in the literature; thus, new synthetic
efforts are urgently required to address this important problem. This task is addressed in the
present investigation, for the first time, by the design and development of novel hydroxyl
functionalized polyesters from L-amino acid resources. This concept was developed by
cleverly combining the solvent-free melt polycondensation route earlier developed from our
laboratory44-52 along with the acetal-masking strategy59-66 in L-aspartic acid system. Acetal-
masking approach has been explored for hydroxyl-functionalized polycaprolactone,59
polylactides,60-61 polyamide, polyurethanes
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63-64,
and sugar-based polyester65-66; however, up
to our knowledge, it is not explored for L-amino acid polymers to make functional polymers.
To accomplish the above task, new multi-functional monomer based on L-aspartic acid was
designed and it was subjected to melt polycondensation with diols to make new classes of
acetal-functionalized L-aspartic polyesters. Post polymerization de-protection of acetal-
polyester yielded new hydroxyl functionalized amphiphilic polyesters. The hydroxyl
polyester geometry was found to exhibit excellent capability for the encapsulation of wide
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ranges of water-soluble and water-insoluble anticancer drugs and fluorophore dyes. The
polyester nano-assemblies are enzyme-biodegradable and this new drug delivery concept is
schematically shown in Figure 1.
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Figure 1. Design and development of novel hydroxyl functionalized polyester based on L-
amino acid bio-resources and demonstration of their enzyme-responsive drug-delivering
capabilities to cancer cells.
Here, hydroxyl functionalized biodegradable amphiphilic polyesters were made
through solvent-free melt polycondensation approach and demonstrated their self-assembled
nano-scaffold as an enzyme-responsive carrier for delivering multiple drugs to the
intracellular compartments of cancer cells. To accomplish this task, the following design
parameters were incorporated: (i) multi-functional L-aspartic acid monomer was designed by
masking the amine into cyclic-acetal moiety and converting the di-carboxylic acids into di-
esters, (ii) under the melt polycondensation process, the acetal unit was found to be inert and
it did not interfere during the transesterification of di-ester with commercial diols, (iii) the
resultant acetal-containing polyester was de-protected to yield hydroxyl functionalized
polyester in which each repeating unit bore bis-hydroxyl functionality to bring hydrophobic-
hydrophilic balance for amphiphilicity in the resultant polymer structure, (iv) the amphiphilic
polymer self-assembled into 250 nm nanoparticle in aqueous medium and exhibited
encapsulation of both water-soluble and water-insoluble anticancer drugs or dyes in a single
nano-carrier platform which is very rarely found in L-amino acid platform. The structures of
the polymers and occurrence of the melt polycondensation process were established.
Dynamic light scattering and atomic force microscope were employed to study the self-
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assembly process. Anticancer drugs doxorubicin, camptothecin, and anti-inflammatory drug-
curcumin, water-insoluble fluorophore-Nile red and congo red, and water-soluble
fluorophore- Rose Bengal were successfully encapsulated in the hydroxyl functionalized
nano-scaffold. Detail photophysical studies like absorption and emission were carried out to
trace the fluorescence properties of the drug/dye loaded nano-scaffolds in the entire visible
light region. In vitro drug release kinetics supported the enzymatic-degradation and enabled
the delivery of cargoes exclusively at the intracellular compartments of the cancer cells. The
cytotoxicity studies proved that the nascent polymers are non-toxic to cells whereas its drug-
loaded nano-carriers accomplished more than 90 % killing in cervical cancer cell lines.
Confocal microscopy studies confirmed the cellular uptake of the drug-loaded nano-carriers
and depending upon the fluorescence properties of the dye/drug-loaded nano-systems the
cancer cell bio-imaging could be accomplished in the entire visible region. The above
investigation proved that the new hydroxyl functionalized polyester is very robust for
accomplishing both therapeutics (by delivering drugs) and diagnostics (via fluorophores
assisted bio-imaging) in L-amino acid bio-resource polymer nano-platform.
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Experimental Section
Materials: L-Aspartic acid, 2,2- bishydroxymethyl propionic acid, dimethoxy propane, 1,12-
dodecanediol, 1,8-octanediol, 1,10-decanediol, p-toulene sulphonic acid (pTSA), triethyl
amine, N’- ethylcarboiimide hydrochloride, titanium tetrabutoxide (Ti(OBu) ),,
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hydrobenzotriazole, di-isopropyl ethyl amine, silicon wafers and pyrene were purchased from
Sigma-Merck Cehmicals and used as it is. Chemicals for In-vitro Bio experiements: Horse
liver esterase enzyme, porcine liver esterase, -chymotrypsin from bovine pancrease,
doxorubicin HCl, curcumin, camptothecin, rose bengal, Nile red, congo red, tetrazolium salt:
3
-(4,5-dimethylthiazolyl)-2,5-diphenyltetrazolium bromide) MTT, glycerol, DAPI (4’, 6-
diamidino-2-phenylindole), Phalloidin–Tetramethylrhodamine isothiocyanate,
B
paraformaldehyde , were purchased from Sigma - Merck chemicals. Trypsin phosphate
versene glucose (TPVG) solution was obtained from Hi-media. HeLa (Cervical cancer cells)
and wild type mouse embryonic fibroblast (WT-MEF) were maintained in phenol red
containing Gibco’s DMEM medium (Dulbecco’s modified eagle medium) with 1% (v/v)
penicillin-streptomycin, 10% (v/v) fetal bovine serum (FBS). 96 and 6-well plastic plates
with flat bottomed were obtained from Costar. Trypsin Phosphate Versene glucose was
obtained from Himedia. Borosil glass based microslides were used for imaging, and
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coverslips were obtained from cornings and rinsed with ethanol and autoclaved prior to use.
Nunc Lab Tek’s 4 well Lab TEK cover glass chamber was used for live cell imaging.
Lysotracker® Green DND-26 was bought from Cell Signalling Technologies (CST). Locally
purchased chemicals: Solvents such as thionyl chloride, DMSO were purchased locally and
used as it is. Methanol was purchased locally and dried and distilled prior to use. AR grade
acetone and DCM were obtained from Finar Chemicals. HPLC THF and HPLC DMSO were
obtained from spectrochem laboratories.
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Methods: H and 13C-NMR were recorded in 400 MHz and 600 MHz Bruker NMR
spectrophotometer. High resolution mass spectrometry-electrospray ionization-quantitative
time-of-flight liquid chromatography-mass spectrometry (HRMS-ESI-Q-TOF LC-MS,
Waters).Mass of polymer aliquots were recorded using Applied Bio system 4800 PLUS
matrix-assisted laser desorption/ionization (MALDI) TOF/TOF analyzer. Thermal
gravimetric analysis (TGA) and differential scanning calorimetry (DSC) of polymers were
carried out using PerkinElmer thermal analyzer STA 600 model and TA Q20 analyzer
respectively. The heating rate and the cooling rate of 10C/min were maintained during the
data acquisition and indium standards were used to calibrate the system. Gel permeation
chromatographic (GPC) plots were determined using Viscotek VE 1122 pump, Viscotek VE
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580 RI detector, and UV/vis detector with polystyrene standards. For Dynamic light
scattering (DLS) sizes of the nanoparticles, Malvern instrument with a Nano ZS-90 apparatus
with inbuild 633 nm red laser (90° angles) was used. The Atomic force microscopy (AFM)
images were generated by VeecoNanoscope IV instrument in tapping mode. TEM samples
were prepared by drop-casting the polymer nanoparticles aqueous solutions on sputter-coated
holey carbon copper TEM grids. These dried samples were analyzed using Technai-300
instrument. PerkinElmer Lambda 45 UV-vis spectrophotometer and SPEX Fluorolog
HORIBA JOBIN VYON fluorescence spectrophotometer were used for all photophysics
analysis of the loaded samples. MTT absorbance readings were recorded using varioskan
FLASH instrument at 570 nm. Christ Alpha 1-2 LD plus was employed to lyophilized
frozen samples. The fixed cell and live cells images were procured using 405, 488, 561 nm
lasers of LSM710 confocal microscope. WT-MEF cells live cell data was procured by
multiphoton laser system (Verdi/Mira 900; Coherent) using the lasers 488 and 561 nm. The
images were acquired using 40 X oil immersion objectives.
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Synthesis of 2,2,5-trimethyl-1,3-dioxane-5-carboxylic acid (1): This compound was
synthesized following the reported procedure.(67) 2,2- Bishydroxymethyl propionic acid (10.0
g, 0.075 mol) and p-toluene sulfonic acid (p-TSA) (0.70 g, 0.004 mol) were taken in a two
neck round bottom flask in dry acetone (100.0 mL). Dimethoxy propane (18.60 mL, 0.112
mol) was added dropwise to the above solution and the reaction mixture was stirred for 4 h at
25 C. Triethylamine (2 drops) was added to neutralize the excess pTSA and the acetone was
removed by rota-evaporation. The product was extracted in ethyl acetate. It was dried over
anhydrous Na SO and the solvent was removed to get crystalline powder as product. Yield =
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0.0 g (77 %). H-NMR in CDCl (400 MHz) ppm: 4.23- 4.20 (2H, d, -CH ), 3.72- 3.69 (
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2H, d, -CH ), 1.47- 1.44 (6H,d, -C(CH )), 1.23 (3H, s, -C(CH )). FTIR (cm ): 3428, 2988,
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880, 1712, 1563, 1456, 1377, 1327, 1246, 1199, 1171, 1155, 1122, 1074, 1035, 988, 933,
03, 829, 761, 729, 716 and 666.
Synthesis of dimethyl (2,2,5- trimethyl-1,3-dioxane-5-carbonyl) aspartate (2): Amine salt
of L-aspartic di-ester was synthesized as reported earlier.50,54 Diisopropylethyl amine (26.5
mL, 0.15 mol) was added into the amine salt (10.0 g, 0.05 mol) in dry dichloromethane
(
DCM) (150.0 mL) and stirred at 25 C until it dissolves completely. Hydroxybenzotriazole
(6.86g, 0.050 mol), N’-ethylcarboiimide hydrochloride (8.6 g, 0.056 mol) and compound (1)
7.06 g, 0.040 mol) were added to the above solution and the reaction was carried out by
(
stirring at 25 C for 30 h. The DCM was rota-evaporated from the reaction mixture and crude
product was extracted in ethyl acetate. The resultant product was purified using silica column
in 30 % ethyl acetate in pet ether. The solvent was evaporated to get pale yellow liquid as a
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product. Yield =16.0 g (75 %). H-NMR in CDCl (400 MHz) ppm : 7.97 (bs, 1H, -NH),
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.83 (s, 1H, -CH), 3.88- 3.76 (dd, 2H, -CH ), 3.70- 3.76 (s, 5H, -CH + -OCH ), 3.59 (s, 3H,
2 2 3
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OCH ), 2.95- 2.78 (dd, 2H, -NH-CH ), 1.37 (s, 6H, -C(CH ) , 0.91 (s. 3H, -C-CH ). C-
3 2 3 2 3
NMR in CDCl (100 MHz): 174.7, 171.2, 98.5. 66.9, 52.6, 51.9, 48.6, 40.1, 36.3, 28.3, 18.4,
3
-1
and 17.6. FT-IR (cm ): 3357, 2992, 2954, 2875, 1735, 1662, 1520, 1438, 1393, 1372, 1346,
1
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315, 1266, 1199, 1178, 1153, 1113, 1079, 1049, 1035, 993, 936, 915, 871, 826, 780, 749,
30 and 689.
Synthesis of acetal-functionalized polyester: Typical procedure for the polymerization
reaction was explained for P12A polymer by melt condensation of monomer 2 with 1,12-
dodecane diol. Monomer 2 (0.62 g, 1.89 mmol) and 1,12-dodecanediol (0.38 g, 1.89 mmol)
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were taken in a melt polymerization tube and the mixture was melted at 100C with constant
48,50
N purging.
2
Ti(OBu) (1.0 mol %) was added as a catalyst to the melt and the tube was
4
degassed at 0.01 mbar vacuum followed by purging with N for five 5 minutes. The
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degassing and nitrogen purging was repeated 3 times. The polymerization was proceeded by
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stirring the melt to 150C for 4h under a N purge. During this process, methanol was
2
removed along with the purge gas and the molten liquid gradually turned into a viscous mass.
-2
The polymerization was continued under a vacuum of 10 mbar for 2 h to produce polymer.
1
Yield= 0.80 g (95 %). H-NMR (400 MHz , CDCl ) δ ppm: 8.04 (s, 1H, -NH), 4.91 (m, 1H, -
3
CH), 4.17 (t, 2H, -COOCH ), 4.09 (t, 2H, -NHCOOCH ), 3.96 (d, 1H, -CH ), 3.88 (d,1H, -
2
2
2
CH ), 3.77 (d, 2H, -CH ), 3.06-2.87 (m, 2H,-CH-CH ), 1.64 ( m, 4H, -COOCH CH ), 1.48
2
2
2
2
2
1
3
(
s, 6H, -C(CH ) ), 1.26 (bs, 16 H, -aliphatic protons), 1.02 (s, 3H, -CH ). C-NMR (100
3
2
3
MHz, CDCl ) δ ppm: 174.1, 171.7, 97.8, 66.9, 65.8, 64.6, 48.3, 39.9, 36.4, 29.4, 28.4, 24.8,
3
-1
1
8.1, and 17.7. FT-IR (cm ): 3324, 2926, 2855, 1733, 1646, 1530, 1461, 1396, 1350, 1187,
1046, 992 and 911.
A similar procedure was followed for the polycondensation of monomer 2 with 1,10-
decanediol and 1,8-octanediol to yield polymers P10A and P8A, respectively, and these
details are given in the supporting information.
Synthesis of hydroxyl-functionalized polyester: The de-protection procedure was explained
in detail for polymer P12H. The polymer P12A (0.46 mg, 1.0 mmol) was taken in a round
bottom flask and dissolved in dichloromethane (DCM) (4.0 mL). Trifluoroacetic acid (0.78
mL, 10.0 mmol), and water (0.80 mL) were added under ice-cold conditions, brought to 25

C, and the stirring was continued for 2 h. The excess solvent was removed from the reaction
mixture using rota-evaporator. The polymer was obtained as a viscous solid mass. Yield=
1
0.39 g (94 %) H-NMR (400 MHz , CDCl ) δ ppm: 8.04 (s, 1H, -NH), 4.91 (m, 1H, -CH),
3
4
3
.17 (t, 2H, -COOCH ), 4.09 (t, 2H, -NHCOOCH ), 3.96 (d, 1H, -CH ), 3.88 (d,1H, -CH ),
2 2 2 2
.77 (d, 2H, -CH ), 3.06-2.87 (m, 2H,-CH-CH ), 1.64 ( m, 4H, -COOCH CH ), 1.48 (s, 6H, -
2
2
2
2
13
C(CH ) ), 1.26 (bs, 16 H, -aliphatic protons), 1.02 (s, 3H, -CH ). C-NMR (150 MHz,
3
2
3
CDCl ) δ ppm:176.7, 171.4, 171.3, 70.7, 68.9, 68.4, 66.5, 65.7, 63.3, 53.8, 48.9, 48.1, 47.2,
3
4
2
1
5.9, 42.2, 41.1, 36.2, 35.9, 32.9, 32.7, 29.8, 29.7, 29.6, 29.6, 29.5, 29.4, 29.3, 28.64, 28.6,
-1
6.0, 25.9, and 25.8. FT-IR (cm ): 3332, 2926, 2654, 1735, 1649, 1533, 1460, 1396, 1350,
281, 1199, 1051, 999, 921, 629 and 608.
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A similar procedure was followed for the de-protection of P10A and P8A to yield
hydroxyl polymers P10H and P8H, respectively, and these details are given in the supporting
information.
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Self-assembly and Encapsulation Studies: The self-assembly of hydroxyl functionalized
polyesters were carried out using the dialysis method explained. About 5.0 mg of the polymer
sample was dissolved in 2.0 mL of DMSO+water (2/3 v/v) and the solution was dialyzed
against water using the semi-permeable membrane of MWCO 1000 Da for more than 48 h.
During this process, freshwater was replenished regularly in the reservoir to remove DMSO.
For loading of drugs, for example doxorubicin loading, 5.0 mg of polymer 0.5 mg
doxorubicin HCl were used. Prior to use doxorubicin HCl was neutralized using tri-ethyl
amine. The dialysis using semi-permeable dialysis membranes of cut off of 1.0 kD ensured
the removal of the un-encapsulated drug molecules. The resultant dialyzed solutions were
filtered, lyophilized and stored at 4C. The lyophilized nanoparticles were re-dispersed in
media prior to use. A similar protocol was followed for the encapsulation of curcumin
(CUR), Camptothecin (CPT), Nile red (NR), Rose Bengal (RB) and Congo Red (CR). For
loading of NR, 100.0 μg was taken as their feed for loading. For photo-physical experiments
and DLS measurements, the dialyzed filtered solutions were used as it is. DLC (Drug
Loading Content) and DLE (Drug Loading Efficiencies) were measured using the equations
1) and (2) as mentioned below:50,54
(
Drug/Dye Loading Content (DLC, in %) =
Dye/ Dye Loading Efficiency (DLE, in %) =
 100 ………..(1)
 100…………(2)
For HR-TEM, AFM images 0.1 mg/mL freshly dialyzed samples were used. The
samples were sonicated for ten minutes prior to drop-casting. Critical aggregation
concentration of the polymer sample was determined using pyrene as a fluorescence probe for
various concentrations of polymer nanoparticles and the procedure has been adopted from the
details from our earlier reports.51
Enzymatic-biodegradation Studies: The biodegradation studies were carried out by taking
the P12H-DOX nanoparticles in a dialysis tube of 1kD cut off and placed in a beaker
containing PBS buffer. The concentration of DOX was taken as 80.0 μg in the total solution.
The solution was incubated at 37 C and the absorbance of the solution outside the dialysis
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membrane was measured at fixed time intervals. To mimic the lysosomal conditions, the
same set up was used, with an additional 10 U of horse liver esterase enzyme, -chemotrysin
from bovine pancreas, porcine liver esterase enzyme and acidic pH 4.0, to study the stability
of these nanoparticles . The absorbance of the solution in the reservoir was measured and the
relative amount of Doxorubicin w.r.t the total initial DOX concentration was plotted against
time as the release profile of the nanoparticles.
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Photophysical studies: The drugs DOX, CPT, and CUR, as well as the dyes NR, RB, and
CR, were loaded in the P12H nanoparticles and were subjected to absorbance and
fluorescence studies to determine the photophysical characteristics of the polymer loaded
nanoparticles. The fluorescence of the drugs and dyes were measured at their respective
absorbance maxima, keeping the slit width 2nm for all the measurements.
Cytotoxicity Studies: Cytotoxicity study was done following our earlier reported procedure
by MTT-assay.50,55 This colorimetry assay was employed to determine the presence of living
or metabolically active cells in the system. The ability of NAD(P)H assisted cellular
oxidoreductase enzyme to reduce the 3−(4,5-dimethylthiazol-2-yl)-2,5- phenyl tetrazolium
bromide (MTT) to purple formazan crystals was employed to determine the number of living
cells present in the system after exposure to the polymers and polymer loaded samples for 48
h. HeLa cell lines ( cancerous) and WT-MEF (Wild type mouse embryonic fibroblast) cell
lines (normal healthy cells) were chosen for this purpose and 1000 cells in each well in a 96
well plate and grown for 24h in a 5 % CO incubator at 37 °C. Various concentrations of
2
P12H-DOX nanoparticles were added and the cells were incubated for 48 h. A control of
viable cells was also maintained and all these experiments were carried out in triplicates. After
48 h and experiment completion, the media was aspirated, followed by addition of 100 μL of a
freshly prepared MTT solution (50.0 μg /mL in DMEM) was added to each well. The treated
cells were further incubated for 4 h at 37 °C under a CO environment. The formazan crystal
2
obtained precipitated out at the bottom of the flat plate due to their insolubility in media. The
media was carefully removed and replaced with 100 μL of HPLC DMSO. The plate was
shaken gently to dissolve the crystals. The photometric absorbance of this purple solution was
collected at 570 nm and average of the three absorbance intensity was used for calculations.
The control cells (without any polymer or samples) were set as standard and the numbers of
metabolically active cells (viable cells) were calculated with respect to these numbers of cells.
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Out of three, if any value was out of standard deviation, then it was omitted during the
calculations. Cell viability percentage was plotted against the concentration of the polymers.
The MTT assays in WT-MEF (Wild type mouse embryonic fibroblast) were carried out for
7
2h following the same procedure.
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Confocal Imaging: This study was done following our earlier reported procedure. 50-52, 54-55
Cornings coverslips were thoroughly washed with 70% ethanol, flame dried and autoclaved
and used for the growing cells. One coverslip each was placed in a 6-well plate and was
washed once with 1.0 mL PBS and media. HeLa cells (passage number 40 and 41 were used
for these experiments). 1 × 105 HeLa cells were seeded in each well in 2.0 mL media and
were grown for 24h under a CO environment at 37 ºC. For bio-imaging applications, freshly
2
dialyzed P12H nanoparticles (loaded with drugs and dyes) were lyophilized for 24 h and were
re-dispersed in media. Concentration of drugs and dyes were taken as 5.0 – 20.0 µM. Cells
were incubated for 5 h and then washed with PBS (1.0 mL) twice to remove all the traces of
the media or suspended fluorophores and drugs that were not taken up by the cells. Cells were
fixed using 4% paraformaldehyde (PFA) solution (in PBS) and stained with DAPI for 2
minutes (for DOX) and were fixed on clean glass slides using mounting media for cell
imaging. In fixed cell imaging for P12H-DOX the cellular distribution was monitored using
phalloidin green to stain the actin filaments of the cells green. The cells were imaged on
confocal microscopy and the DAPI was used for staining nuclei of the cells. The phalloidin
was imaged using 488 nm laser and emission was collected in green channel. DAPI and CPT
were excited at 405 nm laser, and emission was collected at 415- 600 nm (blue channel).
Laser power, gain and airy units for DAPI were 2, 669, and 1.0 A. U, and for CPT was 12,
837, 1.0 A.U. respectively. DOX and CR chromophore was excited at 488 nm and its
emission was collected at 535- 797 nm (red channel), with laser power 3.0, gain 960 and 1.0
A.U. NR and RB were excited at 514 and 561 laser respectively and emission was collected at
red channels. The confocal images were processed in image J software and were processed
using windows and levels in the software. Three regions of interests were selected in the
similar-looking cells and in the background and the corrected total cell fluorescence (CTCF)
calculated following: CTCF = Integrated Density – Mean of the background × area of the cell.
The average of the three values and standard deviations were calculated, and this differential
uptake of the cells was plotted.
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Lysosomal Tracking experiments: A chamber was employed to carry out live-cell imaging
experiments.54 25000 HeLa cells with passage number 40 were seeded a 4 well live-cell
chamber and 300.0 μL media was added and the cells were grown by incubation as mentioned
for cellular imaging for 24h. P12H-DOX nanoparticles were subjected to the cells keeping the
concentration of DOX fixed at 20.0 µM for each experiment. The nanoparticles were added at
different time points to the independent wells at time 5, 15, 60 and 600 min. Free DOX at the
same concentration was also incubated with the cells as a control. After the experiment
completion, the media containing the suspended nanoparticles were aspirated and 300.0 μL of
freshly prepared solution of Lysotracker Green DND-26 (0.15 μL in 3.0 mL media) was
added to each well and the chamber imaged. The Lysotracker Green DND-26 was excited by
laser 488nm and DOX was excited by 561 nm laser and images were collected at green and
red channel respectively. Similar experiments were carried out for WT-MEF cells (passage
21-22). The images thus obtained were processed in Image J software using the windows and
levels tool and were plotted. The CTCF (corrected total cell fluorescence) plots were
generated using Image J software for 8-bit images and a set of 3; three regions of interests
were chosen from the images to calculate the intensity.
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Results and Discussion
Synthesis of Hydroxyl Functionalized Polyesters
L-Aspartic acid bio-resource was selected to make the new classes of hydroxyl
functionalized polyesters by melt polycondensation route as shown in Scheme-1. The
hydroxyl groups in 2,2- bishydroxymethyl propionic acid was reacted with
dimethoxypropane to make cyclic-acetal molecule (1). The reaction of compound 1 with the
amine salt of the L-aspartic di-ester yielded the multifunctional monomer (2) and its structure
was confirmed by NMR spectroscopy in Figure S1. The thermogravimetric analysis proved
that the monomer has very good thermal-stability up to 200 C and confirmed its suitability
for melt polycondensation reaction (Figure S1). Three commercial diols 1,12-dodecanol,
1
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1
,10-decane diol, and 1,8-octane diol were chosen and Ti(OBu) (1 mole % ) was employed
4
as a catalyst for the transesterification polycondensation process. The di-carboxylic esters in
the monomer reacted with diols to yield aliphatic ester chemical linkages under the melt
transesterification process while the amide and acetal functional units in the pendants were
inert during this process. The polymerization was carried out first under nitrogen purge for 4
h at 150 C and then under vacuum (0.01 mbar) to produce polymers. The resultant aliphatic
polyesters have one cyclic-acetal functional group in each repeating unit and these polymers
are referred to as P12A, P10A, and P8A with respect to the number of carbon atoms in the
diol segment and “A” for acetal groups (see Scheme-1).
OCH3
O
HO
HO
O
OCH3
O
O
O
OH
O
O
HO
(i) Ti(OBu)4, 150 C, 4 h
ii)vacuum, 2 h
OH
OH
x
PTSA
EDC.HCl
DIPEA
O
1
0
Acetone
O
HN
O
O
HN
O
O
Dry DCM
24 h, 30 0
C
(
O
OCH3
+
NH3 Cl-
O
NH2
O
O
H3CO
x
n
SOCl2
MeOH
O
2
O
OH
OCH3
HO
H3CO
P8A (x=4); P10A (x= 6) and P12A (x=8)
O
L-Aspartic acid
Acetal-functionalized L-Aspartic Polyester
Scheme 1. Synthesis of acetal functionalized polyesters from L-aspartic acid.
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Figure 2. NMR spectra of monomer (a and c) and polymer P12A (b and d) were recorded in
CDCl . The different atoms are assigned by alphabets and the solvent peaks are indicated by
3
asterisks.
1
The H-NMR spectrum of the monomer and the polymer P12A are given in Figure 2
and the different types of protons and carbons are assigned by alphabets. The methyl ester
peaks in the monomer at 3.67 and 3.59 pm (peak a and a’ in Figure 2a) diminished and new
esters peaks with respect to the formation of polymer chains RCOOCH CH appeared at 4.17
2
2
and 4.09 ppm (peaks b and b’ in Figure 2b). The acetal group –CCH OC(CH ) at 1.48 ppm
2
3 2
(
peak c) and multiple peaks at the region of 3.9 to 3.6 ppm with respect to –CCH OC(CH )
2
3 2
(peak d) were found to be completely intact in both monomer and polymer spectra. This
observation confirmed that the acetal unit was completely inert and it was not disturbed
during the melt polycondensation reaction. The comparison of ratios of intensity of new ester
peaks RCOOCH CH (peaks at 4.17-4.09 ppm, b and b’) with that of the methyl esters at the
2
2
chain ends (peaks at 3.67-3.59 ppm) in the polymer spectrum enabled the determination of
the number average repeating units (X ) in the polymer chains. The X was determined as 30-
n
n
3
2 in P12A and the substitution of this value in Carothers equation [X = 1/(1-p)] gave the %
n
of reaction, p > 98 %. The occurrence of the polycondensation process was further supported
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by C-NMR (Figures 2c and 2d). The peaks of major importance were the methyl ester
carbon peaks at 52.6 and 51.9 ppm in the monomer (Figure 2c) which completely
disappeared and new ester carbon peaks appeared in the polymer spectrum at 65.7 and 65.0
ppm (see Figure 2d). Interestingly the acetal carbon atoms -CCH OC(CH ) and -
2
3 2
1
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CCH OC(CH ) at 98.5 ppm and 17.7, respectively, in the polymer and monomer spectra
2
3 2
were not disturbed. Based on the above analysis, it may be concluded that (i) the
multifunctional L-aspartic acid monomer 2 was stable under melt condition to condense with
commercial diols to produce linear acetal-containing polyesters. Similarly, the structures of
the polymers P10A and P8A were confirmed by NMR and the details are given in Figure S2.
The molecular weights of the polymers were determined by GPC in tetrahydrofuran solvent
and their respective chromatograms are shown in Figure 3a and the molecular weights were
summarized in the table in Figure 3b. The P12A acetal-polymer showed the formation of
3
3
high molecular weights as M = 12x10 g/mol and M = 25x10 g/mol with a polydispersity
n
w
of 2.3. The P10A and P8A polymers showed the formation of moderate molecular weights in
3
3
the range of M = 5x10 g/mol and M = 18x10 g/mol.
n
w
Figure 3. (a) GPC chromatograms of acetal functionalized polyesters in tetrahydrofuran at
5 C. (b) GPC molecular weights and thermal properties of the polymers are shown in the
table. (c) GPC chromatograms of polymer P12A aliquots taken at various time intervals. (d)
MALDI-TOF mass spectrum of P12A aliquots taken at 2 h. The inset table shows the
calculated and found mass peaks for different types of chain ends for trimer species.
2
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To further study the molecular weight build-up in the polycondensation reaction;
polymerization kinetic was carried out for the P12A polymer synthesis. Polymer sample
aliquots were taken at regular time interval and they were subjected to NMR and GPC
analysis. The GPC plot of the aliquots in Figure 3c revealed the formation of higher
molecular weight chains with an increase in reaction time. The GPC plots gradually changed
from multi-model to narrow disperse samples as a result of the transformation from
oligomers to high molecular weight polymer chains in the condensation process. The NMR
spectra of the aliquots in Figure S3 showed the disappearance of the methoxy protons and
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appearance of the new ester peaks as observed in Figure 2. The plot of M versus %
n
conversion showed typical trend for condensation polymerization with respect to increase in
the molecular weight at higher % conversion (see Figure S4). The polydispersity of the
aliquots were found to be ~ 2.0 which is typically expected for condensation polymers. To
study the non-interference of the acetal units during the polycondensation process, the
aliquots were subjected to MALDI-TOF MS analysis for end group analysis. A MALDI-TOF
spectrum for 2h aliquot is shown in Figure 3d and the peaks are assigned for different types
of chain ends (for MALDI-TOF spectra of other aliquots, see Figure S5). In a typical A-A +
B-B polycondensation, the growing chains are expected to have four types of reactive species
th
such as A-P -A, A-P -B, B-P -B and P -cyclic, where P represents the repeating unit of ‘n ’
n
n
n
n
n
chain. In the present case, each repeating unit mass is corresponding to 455.48 g/mol; thus,
for the trimer species, the mass for the different types of chain ends are calculated and shown
in the table inset. The peaks in the MALDI-TOF spectrum are assigned with respect to these
chains showed in Figure 3d. Two important points are clear from these studies; (i) the peaks
in the MALDI-TOF spectrum preferably matched with the expected repeating unit mass
having protected acetal chemical linkages, and (ii) there are no traces of the peaks with
respect to the macro-cyclic (P ) formation. These observations confirmed that the acetal units
n
were stable under melt polycondensation process and also the polymerization preferably
underwent via linear chain formation to produce higher molecular weight chains. This
revealed that the melt transesterification process applied for L-aspartic acid monomer is
stable to produce the acetal-functionalized polyesters under solvent-free process.
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O
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OH OH
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CH Cl2
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n
O
O
P8A (x=4); P10A (x= 6) and P12A (x=8)
P8H (x=4); P10H (x= 6) and P12H (x=8)
Scheme 2: Synthesis of hydroxyl functionalized L-aspartic acid based polyesters.
The de-protection of the above acetal-polyesters yielded their corresponding hydroxyl
functionalized polyesters P12H, P10H, and P8H (H-hydroxyl) (see Scheme-2). For this
purpose, the acetal-polymers were subjected to de-protection under acidic medium (using
trifluoroacetic acid, TFA) to yield the hydroxyl functionalized polyesters as shown in
Scheme-2. The complete de-protection of the acetal units without affecting the polyester
1
backbone was confirmed by NMR spectroscopy analysis. H-NMR spectra of the hydroxyl
functionalized polymers are shown in Figure S6. The polymer spectra showed the
disappearance of the acetal units (vanishing of peak c at 1.48 ppm) and appearance of the
new hydroxyl units R-CH OH at 3.8-3.5 ppm in the polymer (see Figure S6). Further, the
2
comparisons of the peak intensities in the hydroxyl polyester in Figure S6 with respect to the
ester peaks RCOOCH CH (peaks at 4.11-4.19 ppm) and hydroxyl units R-CH OH at 3.8-3.6
2
2
2
ppm matched very well with 1:1 ratio for the expected polymer structure. The de-protected
polymer sample was subjected to HRMS analysis and the data is shown in the supporting
information Figure S7. The deprotected polymer samples showed peaks with respect to the
expected hydroxyl functionalized polymer structure. The peaks corresponding to the A-P -B
n
and B-P -B where n= 1-3 were clearly visible which further supported the complete de-
n
protection of acetal to hydroxyl functional groups. This confirmed that the de-protection was
accomplished without affecting the aliphatic polyester backbone. GPC molecular weights of
the hydroxyl functionalized polymers were determined by GPC and their plots are shown in
Figure S8 and the molecular weights are summarized in Table 3b. The hydroxyl
functionalized polymer exhibited higher retention time compared to its acetal-polymer and
this trend was attributed to the variation in the hydrodynamic volume of these two different
polymer structures in tetrahydrofuran (GPC solvent). The GPC plots of P12H and P10H
could be clearly visualized whereas the P8H polymer eluted along with the solvent (see figure
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S8). The thermal properties of the polymers were studied by TGA and DSC, and the data are
shown in Figure S9 and S10. The thermal stability of the acetal-polymers was found to be
very good in the range of 290-310 C whereas their hydroxyl functionalized polymers
showed stability only up to 240-250 C. The relatively less thermal stability of the hydroxyl
functional polyesters are not clearly understood at present and it may be partially attributed to
thermal hydrolysis of aliphatic ester backbone by the back-biting of hydroxyl groups.
Nevertheless, this is not a limitation on the hydroxyl polymers for biomedical applications
since they have stability up to 250 C which is sufficient enough for drug delivery
application. DSC thermograms of the polymers showed only glass transition temperature (Tg)
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with respect to amorphous polymers. The T of the acetal-polymers were found to be -15 to -
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higher T values for the hydroxyl polymers is attributed to the possibility for the existence of
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hydrogen bonding interactions among the hydroxyl functional groups at the periphery. Based
on the above analysis, it may be summarized: (i) the melt transesterification underwent more
than 98 % to produce high molecular weight chains, and (ii) the de-protection of the acetal-
polymers yielded new classes of hydroxyl-functionalized polyesters, for the first time, from
L-aspartic acid in the literature. Hence, the new synthetic strategy is very good to produce
new classes of well-defined acetal-containing and hydroxyl functionalized aliphatic
polyesters from L-amino acid bio-resources. Among all three polymers, the hydroxyl
functionalized polymer synthesized using 1,12-dodecanediol possessed higher molecular
weights; thus, the polymer P12H was further chosen to demonstrate the biomedical
application of these newly designed structures in drug delivery to cancer cells.
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Self-assembly and Drug Encapsulation
The L- aspartic acid based hydroxyl functionalized polymers have unique structural
geometry with hydrophobic polyester backbone and bis-hydroxyl groups in every repeating
unit providing hydrophilic at periphery. As a result, the polymer structures gain appropriate
amphiphilicity to self-assemble in the aqueous medium. The side chain part having the
hydroxyl units tends to phase segregate away from the hydrophobic backbone; thus it
produces hair-pin bend type amphiphiles in the aqueous medium. The subsequent aggregation
of these amphiphiles self-assembled into polymer nanoparticles with hydrophobic core and
hydroxyl groups containing periphery as shown in Figure 4a. To produce the self-assembled
nano-assemblies typically the hydroxyl polymers were dissolved in DMSO+water (2:3 v/v)
solvent mixture and dialyzed using semi-permeable membrane having MWCO of 3500 Da.
The reservoir was replenished periodically with fresh water and this facilitates the complete
removal of the DMSO and other impurities. At the end of the dialysis clear solution was
obtained which was lyophilized and the sample was stored at 4 C. To study the
encapsulation capabilities of the polymer nano-assemblies, a similar dialysis procedure was
followed in which the desired drugs or fluorophore dyes were also taken along with the
polymer. The un-encapsulated drugs or dye molecules were removed continuously by
replenishing with fresh water in the reservoir until otherwise the solution became colorless (at
least for 48 h). Two clinically important anticancer drugs doxorubicin (DOX) and
camptothecin (CPT), and anti-inflammatory natural drug curcumin (CUR) were chosen.
Three fluorophore dyes (or markers) Nile red (NR), Rose Bengal (RB) and Congo Red CR)
employed routinely for bio-imaging application were also chosen for encapsulation. It is
important to mention that DOX and RB are water-soluble whereas all other four molecules
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(CUR, CPT, NR, and CR) are water-insoluble. Hence, the selections of these drugs/dyes
provide appropriate choice to check the encapsulation capabilities of the newly designed
hydroxyl functionalized polyester P12H. The P12H encapsulated nanoparticles are referred
to as P12H-DOX, P12H-CPT, P12H-CUR, P12H-NR, P12H-RB, and P12H-CR. The
dialyzed polymer solution containing these drugs and dyes in the vial was shown in Figure
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Figure 4: (a) Schematic representation of self-assembly of hydroxyl functionalized L-aspartic
acid nanoparticles and their drug or dye loading capabilities. The photographs showing the
DOX, CPT, CUR, NR, CR, and RB encapsulated vials along with nascent polymer under
normal light and upon photo-excitation. (b) DLS histograms of DOX loaded nanoparticles
P12H-DOX and the schematic representation of aggregated nanoparticles. (c)High-
resolution transmission electron microscope images of P12H-DOX. The figure in the inset
shows the HR-TEM image of the aggregated polymer nanoparticles. (d) Atomic force
microscopy image of P12H-DOX and the height profile of the nanoparticles.
The aqueous solution nano-aggregates of the P12H were studied by dynamic light
scattering (DLS) and the DLS histogram of P12H-DOX is shown in Figure 4b. DLS studies
showed that P12H-DOX exhibited mono-modal histogram with size 255  40 nm. The PDI
of the P12H-NR, P12H-CR, P12H-CUR, P12H-RB, P12H-DOX, P12H-CPT and P12H
nanoparticles were found to be in the range of 0.12 to 0.30. The DOX loaded nanoparticles
were further subjected to high-resolution transmission electron microscope imaging and the
TEM image in Figure 4c showed the formation of 25-30 nm nanoparticles and their
subsequent aggregation into large nanoparticles of 180 nm. High resolution TEM images
clearly showed the co-existence of the aggregation of many nanoparticles together to produce
larger size aggregated nanoparticles of 180 nm in size. During the self-assembly process, the
amphiphilic polymer chains collapsed into coil-like form which typically undergoes
aggregation to produce individual nano-particles of typically 15-20 nm in size. These
nanoparticles further undergo aggregation to produce larger nanoparticles of 180 nm in size
(shown as inset in Figure 4c). The hydroxyl substituted polymers reported here has clear
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hydrophilic (hydroxyl units) and hydrophobic (back bone) domains within their structure,
thus the phase separation facilitated the observation of both individual nanoparticles and their
aggregation together into exhibiting larger nanoparticle size formation (shown by circle) in
the TEM image in Figure 4c. Additional TEM images are given in the supporting information
Figure S11 to support this aggregated nanoparticle formation. DLS histograms typically
showed the hydrodynamic diameter of the aggregated nanoparticles in the aqueous medium;
thus, these nano-aggregated assemblies matched with the aggregated nanoparticles in HR-
TEM image. Atomic force microscope (AFM) is a relatively soft technique to measure the
size of the nano-aggregates which are directly drop casted on the mica surface. AFM image
in Figure 4d of the P-12H-DOX showed the existence of smaller (50 nm) as well as larger
nanoparticles (220 nm). The above analysis suggested that the hydroxyl polyester P12H
nanoparticles self-assembled into smaller nano-assemblies which underwent aggregation to
produce nearly 200 nm nanoparticles. The height and width of the nanoparticles measured by
AFM is showed along with Figure 4d. The half-width at maximum showed the size of the
nanoparticle as 200 nm and supported the size of the nanoparticle measured in HR-TEM and
DLS. DLS histograms of other drug-loaded and fluorophore loaded nanoparticles are shown
in the supporting information in Figure S12. These drug or dye loaded nanoparticles showed
the average sizes < 250 nm and matched with the DLS and TEM observation of P12H-DOX
sample. The critical aggregate concentration (CAC) of the polymer was determined using
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pyrene as chromophores following the literature report . Typically 0.6 M concentration of
pyrene was employed for the CAC studies by varying the polymer concentration ranging
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from 1×10 to 1×10 mg/mL. The I /I ratio of the emission peak intensities in pyrene was
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plotted against the concentration of the polymer for the CAC determination (see Figure S13).
-6
The break point was observed at 0.53×10 g/mL with respect to the CAC of the newly
designed hydroxyl polyester. This further supports that the polymer structure possessed
unique amphiphilic nature to produce self-assembled micellar structure in the aqueous
medium at micro-molar concentration range. The CAC of the drug or dye loaded samples
could not be determined due to the overlap of fluorescence signals of the encapsulated dyes
with pyrene. Thus, the DLS data of the nascent polymer with their dye and drug encapsulated
polymer nanoparticles were compared to indirectly understand the stability. The DLS
histograms of drug loaded polymer are provided in Figure S12 and support that the polymer
nanoparticle size did not change before and after encapsulation of drugs or dyes. The above
studies revealed that the custom-designed hydroxyl functionalized polymer nano-assemblies
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are very robust in encapsulating both water-soluble and water-insoluble drugs and
fluorophore dyes without altering the size and shape of the nanoparticles.
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Figure 5. (a)Emission spectra of the drug and fluorophore encapsulated P12H nanoparticles
(exc = absorbance maxima). (b) The table containing the average size of the drug and
fluorophore encapsulated P12H nanoparticles determined by DLS; drug loading content
(DLC) determined by absorption spectroscopy, drug loading efficiency (DLE), their
absorbance and emission maxima.
The drug molecules DOX, CPT, CUR and the dye molecules RB, NR, CR are highly
luminescent molecules; thus, their absorbance and emission spectra were recorded for the
aqueous nanoparticles. Absorbance spectra are shown in Figure S14 and their emission
spectra are shown in Figure 5a. The DLS size of the nano-aggregates, their absorbance, and
emission maxima values are shown in the table in Figure 5b. The encapsulated fluorophore
dyes are predominantly red-color thus they exhibited absorbance maxima from 500-550 nm
and emission maxima at 560-620 nm. On the other hand CPT, CUR and DOX loaded
samples showed blue, green and red emission, respectively, with distant maxima at 433, 510
and 554 nm. Thus, the present approach provides significant color-tunability in terms of
encapsulated drug or fluorophore dyes ranging from 300 to 650 nm in the entire visible light
region in a single polymer nano-platform. The drug loading content (DLC) and drug loading
efficiency (DLE) seems to be varying depending upon the loaded cargoes. DOX exhibited
higher loading compared to all other molecules. Nevertheless, the number of drugs and
fluorophore dyes are significant enough to study their cytotoxicity effect and bio-imaging in
cell lines. Additionally, the present hydroxyl polymer system enabled the encapsulation of
almost 6 different cargoes; thus, their cellular uptake studies would be valuable in the L-
amino acid-based nano-carriers in the literature.
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Enzymatic-Biodegradation and Cytotoxicity Studies
The polymer backbone is designed with aliphatic ester chemical linkages which are
well known to undergo biodegradable cleavage by lysosomal enzymes at the intracellular
environment. Earlier attempts revealed that esterase enzymes and proteomic enzymes like
trypsin and -chymotrypsin potentially degraded the aliphatic polyesters and aliphatic amide
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chemical linkages polymeric nanoassemblies. To study the enzymatic biodegradation of the
polymer backbone in P12H chains, the DOX loaded nanoparticle was subjected to in vitro
drug release studies in the presence of various enzymes. In vitro drug release study was done
by dialysis method and the drug release content was measured by absorption spectroscopy
and the cumulative drug release was determined. In the absence of exposure to enzymes, the
P12H-DOX nanoparticles experience extracellular like conditions; thus, the degree of the
cleavage of the nanoparticles is expected to be minimum to nil. On the other hand, once the
nanoparticles were taken across the cell membrane, they are readily exposed to the enzymes
at the intracellular level for biodegradation to release the drugs as shown in Figure 6a. In
vitro drug release data for the P12H-DOX nanoparticles in PBS at 37 C (pH 7.4) is shown in
Figure 6b. It was found that the doxorubicin encapsulated nanoparticles was stable in the
nano-cavity under (in PBS) physiological conditions. To study the biodegradation of nano-
scaffold by the lysosomal enzymes, the drug release kinetics was investigated by incubating
the P12H-DOX in the presence of 10 U of three enzymes such as horse liver esterase, porcine
liver esterase and -chymotrypsin from bovine pancreas in PBS at 37 C (pH 7.4). These
enzymes are known to cleave the aliphatic polyesters and aliphatic polyamide linkages which
de-stabilize the loading capabilities of the polymer nanoparticles and released the cargoes.54
Doxorubicin was found to be released nearly 40-50 % within 12 h exposure to these enzymes
which was very significant compared to the release profiles in the absence of the enzymes
(see Figure 6b). Among all three enzymes, -chymotrypsin was found to exhibit more
biodegradation since it has capability to degrade both the polyester backbone and amide side-
chain in P-12H structure. Surprisingly, the complete release of the doxorubicin from the
nano-scaffold was not observed even after 36 h which is not clear at present. This may be
partially due to the strong interaction of the hydroxyl units with DOX which restrict the
enzymatic-biodegradation at large. Further structure-optimization of hydroxyl polyesters
may be required in the long term to accomplish complete release of the drugs and validate
this trend. The stability of the polymer nanoparticles were studied in different pH. The
polymer nanoparticles were incubated at pH=4.0 (acetate buffer) 68 and pH=7.4 (PBS) and
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their stability was studied using dynamic light scattering at 37 C for 24 h (see Figure S15). It
was found that the nanoparticles were stable in the acidic pH. This data further supports the
observation in Figure 6b that the polymer nanoparticle was susceptible to biodegradation
exclusively in the presence of enzymes. Nevertheless, the current design was able to release
about 50 % of the drugs in a controlled manner upon exposure to lysosomal esterase enzymes
which validated the intracellular drug-delivering capacities of the hydroxyl functionalized
polyester nanoparticles.
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Figure 6. (a) Schematic representation of enzymatic-biodegradation of P12H-DOX
nanoparticles at intracellular compartments. (b) Cumulative drug release profiles of P12H-
DOX in the absence and in the presence of horse liver esterase enzyme, -chymotrypsin from
bovine pancreas and porcine liver esterase at 37 C and pH 7.4 in PBS. (c) Live-cell
confocal microscopy imaging of P12H-DOX nanoparticles in the presence of lyso-tracker in
HeLa cell line. The concentration of DOX is maintained as 20 M and incubated for 6h. (d)
Confocal microscopy imaging of variable temperature dependent cellular uptake studies in
HeLa fixed cells. The concentration of DOX is maintained as 20 M and incubated for 5h.(
Scale bar in the cell images= 50m and 25m ).
HeLa cells were chosen to carry out cellular uptake of P12H-DOX nanoparticles to
corroborate the enzyme degradation at cellular levels. Since the lysosomes of cells are rich in
hydrolytic enzymes, this experiment was carried out to establish the uptake of the
nanoparticles via endocytosis. The P12H-DOX nanoparticles were incubated with HeLa cell
in a live cell chamber for 6 h. The free dispersed nanoparticles in media that were not taken
up by the cells, if any, were removed by aspirating the media. To rule out the possibility of
the uptake of the nanoparticles by diffusion or another method, the lysosomes of the cells
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were stained with highly selective Lysotracker Green DND dye. This staining was carried out
immediately before the imaging of the live cells. The Lysotracker Green DND was excited at
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88 nm with laser power 2 ( keeping gain ˂ 800) at 1.0 A.U. and was imaged in the green
channel. The DOX moieties were excited at 561 nm laser and were imaged in the red
channel. Smart set up was used to select the tracks for exciting the fluorophores to eliminate
the cross-talk between the emission and excitation of the Lysotracker Green DND and DOX.
The images thus obtained were processed in Image J software and were plotted and are
shown figure 6c. The four panels show the differential interference contrast (DIC), lyso green
channel, DOX channel and the merged channel (red and green channel merged). The DIC
channel showed the cell position and morphology during the live cell experiment. The green
channel showed the bright green color that emerged from the lysosomal compartments of the
cellular regions. These regions are highly acidic in nature and are selectively stained by the
Lyso-tracker Green DND. The third panel shows the DOX uptake in the HeLa cells in 6 h
and it was observed that DOX emission was obtained from all over the cells with high-
intensity regions in few places. On merging the green and the red channel it was observed
that this high concentration of DOX was indeed present at the lysosomal regions, as the
colocalization of lysosomes and DOX could only produce bright yellow emission (a mixture
of red and green). Therefore, within 6h of incubation, the P12H-DOX successfully
accumulated in the lysosomal compartments of the cell. The P12H nanoparticles were
excellent carriers for DOX and selectively deliver the DOX in the HeLa cells as these would
undergo lysosomal degradation, thereby releasing the drug into the cytoplasm.
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To further support the endocytosis assisted the uptake of the P12H nanoparticles in
the cells a time dependent uptake experiment was carried out for fixed cell imaging. Since
endocytosis is an energy-dependent process hence it is believed that any process occurring
due to endocytosis should slow down at temperatures lower than the physiological
temperatures.69-70 Therefore, a temperature-dependent experiment was carried out wherein,
P12H nanoparticles loaded with DOX, CUR, and CPT were incubated with cells at different
temperatures. HeLa cells were seeded in two separate 6-well plates and grown for 24 h in an
incubator with 5% CO at 37 °C. After 24h the media from the cells were aspirated and 20.0
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μM of DOX, CUR and CPT in P12H-DOX, P12H-CUR, and P12H-CPT nanoparticles (in
media) was added to the cells. One 6-well plate was incubated at 37 °C and the other plate
was maintained at 4° C. The cells were fixed after 5h incubation. The cells in case of DOX
chromophores were stained with DAPI prior to mounting on glass slides. Further, these cells
were imaged. DAPI, CPT, and CUR were excited at 405 nm laser and were imaged in blue,
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blue and green channels respectively (see Figures S16 and Figure S17). The confocal images
of the P12H-DOX at 4°C and 37°C have been shown in figure 6d. The merged image shows
the DAPI stained blue nucleus and the red emission from the DOX distributed in cytoplasm
of the cells. It was clearly evident that at lower temperatures such as 4°C the uptake of
nanoparticles was low compared to the uptake at physiological temperature as red emission
from the cells was observed more in case of 37°C. Further, the CTCF intensities of DOX
were calculated and plotted in figure 6d. It was observed that uptake of DOX at 37°C was 3
times higher compared to that at 4°C. This was evident because at lower temperature
conditions the process of the endocytosis was slowed down and resulted in lesser uptake of
the P12H nanoparticles. This was further extended to P12H-CPT and P12H-CUR
nanoparticles and the images have been shown in Figures S16 and Figure S17. A similar
trend was observed wherein lower emission was observed in the case of P12H nanoparticles
incubated at 4 °C both in the blue (cyan) and green channel for CPT and CUR respectively.
Whereas the cells showed higher uptake of the P12H-CPT and P12H-CUR at 37 °C, further
illustrating the endocytosis dependent uptake of the nanoparticles in the cells. Since the size
of free drugs is very less compared to nanoparticles (~ 5-10 nm), they are taken up by simple
diffusion across the cell membrane. Therefore, it was anticipated that there should not be
much difference in uptake of free drugs at either temperature. The uptake of the free drugs
was found to be similar as seen in the Figure S16 Figure S17. Hence, compared to
temperature-dependent uptake of free drugs, the P12H nanoparticles were taken up more in
the cells and the same was corroborated by the live-cell experiments. It was found that the
P12H nanoparticles accumulated in the lysosomes of the HeLa cells, which was essential for
their biodegradation for the delivery of the loaded cargoes.
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Figure 7: Cell viability of P12H nanoparticles (a), P12H-RB nanoparticles (b), free DOX
and P12H-DOX nanoparticle (c), free CUR and P12H-CUR (d) in HeLa cells line at 37°C
for 48 h. (e) Cell viability of P12H nanoparticles in WT-MEF cell line at 37°C for 48 h. (f)
Cell viability of free DOX and P12H-DOX nanoparticle in WT-MEF cell lines at 37°C for 48
h.
The biocompatibility of nascent polymer P-12H was investigated in cervical cancer
cell lines (HeLa) and wild type mouse embryonic fibroblast (WT-MEF). As can be seen in
Figure 7a, the nascent polymer nanoparticles were not cytotoxic to the HeLa cells even at a
concentration up to 100 µg/mL and were highly suitable as drug delivery carriers. These
polymers exhibited the unique quality of being able to load and stabilize multiple dyes and
drugs. The cytotoxicity profile in HeLa cells of the polymer nanoparticle loaded with Rose
Bengal (P12H-RB) has been shown in Figure 7b, which exhibits that these dye loaded
nanoparticles are highly biocompatible up to a concentration of 100 µg/mL. The polymers
also successfully encapsulated anticancer drugs like Doxorubicin and Curcumin and the
anticancer efficacy of these drug loaded nanoparticles were studied in HeLa cell lines. For the
free DOX and DOX-loaded nanoparticles (P12H-DOX), the concentration of the drug was
varied from 0.1 to 1 µg/mL and then the MTT assay was carried out after 48h of compound
incubation. As can be seen in Figure 7c, the cell growth inhibition capability of the free drug
saturated at a concentration of 0.4 µg/mL and it was able to achieve 50 % cell killing at the
concentration of 1.0 µg/mL. On the other hand, the DOX-loaded nanoparticle was able to
accomplish ~ 80 % cell growth inhibition and the EC value exhibited by these nanoparticles
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was 0.4 µg/mL, in accordance with the literature reports. In the case of Curcumin, the
concentration was ranged from 1 to 75 µg/mL and the MTT assay is shown in figure 7d. The
free Curcumin was able to achieve ~ 80 % cell killing with an EC value of 10 µg/mL
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whereas the polymer loaded curcumin (P12H-CUR) nanoparticles exhibited 50 % cell killing.
Cytotoxicity data for P12H-CPT, P12H-NR, and P12H-CR are shown in the supporting
information Figure S18. Based on the above studies, it may be summarized that the newly
designed P12H polymer is very effective in inducing cell killing for the drug loaded nano-
carriers (DOX, CPT and CUR) whereas their fluorophore (NR, CR, and RB) loaded
nanoparticles are largely non-toxic to HeLa cell lines. In a similar way, P12H-DOX and
P12H nanoparticles were also incubated with non-cancerous healthy cell lines WT-MEF
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(
Wild- Type Mouse Embryonic Fibroblast cells) and the cell viability plots have been shown
in Figures 7e and 7f. It was observed that significant cell killing (~ 80%) was achieved by
P12H-DOX loaded nanoparticles containing 1.0 g/mL DOX, whereas the higher amounts of
the polymer (~100 g/mL) showed extremely low toxicity towards the cells thereby
demonstrating the effective therapeutic efficacy of the L-aspartic acid based newly designed
P12H polyesters.
Cellular Uptake Studies
The ability of the P12H-DOX nanoparticles as drug carrier was demonstrated by
studying the cellular uptake of doxorubicin in HeLa and WT-MEF cell lines. The killing of
cells occur via DNA intercalation mechanism of doxorubicin in the nucleus of the cell,
therefore determination of the location of doxorubicin within the cell was crucial to analyze.
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HeLa cells were grown in a 6 well plate at a concentration of 10 cells in each well. The cells
were further treated with doxorubicin, fixed and were subjected to confocal microscopy
imaging. DAPI was used to stain the nucleus, and Alexa dye conjugated phalloidin green was
used to illuminate cytoskeleton actin filaments network of the cells. This experiment was
conducted to determine the location of the DOX molecules after the lysosomal accumulation
of the nanoparticles in the cells (see figure 8). A control for cells without DOX was also
maintained to rule out the possibility of signal interference from the other chromophores. The
free doxorubicin and P12H-DOX nanoparticles were exposed to the HeLa cells keeping the
concentration of the doxorubicin molecule same, to understand the efficacy of the nano-
particle in drug delivery. The fixed cells were imaged using 405, 561 and 488 nm lasers for
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DAPI, DOX and Phalloidin respectively. The confocal images are shown in figure 8a. The
DIC images showed the fixed cells, and DAPI channel showed the nucleus in the cells, the
DOX channel showed the location of DOX molecules and the Phalloidin channel showed the
actin filaments of the cells. This was further merged in a fifth panel in figure 8a to show the
colocalization of the three fluorophores. The merged image shown in figure 8a showed clear
demarcation of the nucleus as well as doxorubicin uptake in the cells. Free DOX diffused in
the cells and was found to accumulate in the nucleus and was shown by the magenta color
because of the overlap of the DAPI and the DOX. The nanoparticles, however, were taken up
by the endocytosis and the DOX was delivered to the cytoplasm of the cells and it was found
to accumulate in the nucleus of the cells as shown in figure 8a. It was interesting to note that
the emission intensity of DOX from the nucleus was higher than the free DOX and it was a
qualitative-evidence towards higher uptake of the DOX in the cells. Therefore, the CTCF
calculations were carried out to quantify it and the plots have been shown in figure 8b. It was
observed that P12H-DOX carried ten folds higher DOX drug into the cells which is superior
compared to simple diffusion of free DOX across the cell membrane. Hence, the P12H-DOX
nanoparticles were capable of delivering DOX in to the cell, which would cause cell killing
which was reflected in the cytotoxicity data as shown in figure 7c. Doxorubicin is a small
molecular drug with size 50 Å; thus, it is known to efflux out from the cells through
membrane efflux pumps. On the other hand, the DOX loaded nanoparticles are significantly
larger in size; hence they localized in the cytosol for longer period which enhances the
probability of the DOX interaction with nucleus for increased cytotoxicity effect. Further, the
live cell cellular uptake studies clearly evident for the enhanced uptake of the DOX (see
Figure 6c) while delivered from the newly designed hydroxyl polymer P12H platform
compared to the free drug. Thus, both the nanoparticle design and the increased cellular
uptake enhanced the anticancer activity of P12H-DOX nanoparticle.
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