Alginate Bead Based Hexagonal Close Packed 3D Implant for Bone

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Alginate bead based hexagonal close packed 3D implant for bone tissue engineering Tarun Agarwal, Prajna Kabiraj, Gautham Hari Narayana, Senthilguru Kulanthaivel, Uvanesh Kasiviswanathan, Kunal Pal, Supratim Giri, Tapas Kumar Maiti, and Indranil Banerjee ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.6b08512 • Publication Date (Web): 07 Nov 2016 Downloaded from http://pubs.acs.org on November 12, 2016

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ACS Applied Materials & Interfaces

Alginate bead based hexagonal close packed 3D implant for bone tissue engineering Tarun Agarwal a, c, Prajna Kabiraj a, Gautham Hari Narayana a, Senthilguru Kulanthaivel a, Uvanesh Kasiviswanathan a, Kunal Pal a, Supratim Giri b, Tapas K. Maiti c, Indranil Banerjee a* a Department of Biotechnology and Medical Engineering, b Department of Chemistry, National Institute of Technology, Rourkela, Odisha, Pin: 769008, India c Department of Biotechnology, Indian Institute of Technology Kharagpur West Bengal, Pin: 721302, India Mr.Tarun Agarwal Department of Biotechnology and Medical Engineering, National Institute of Technology, Rourkela Odisha, Pin: 769008, India Email: [email protected] Phone: +919933968910 Ms. Prajna Kabiraj Department of Biotechnology and Medical Engineering, National Institute of Technology, Rourkela Odisha, Pin: 769008, India Email: [email protected] Phone: +919819835621 Mr.Gautham Hari Narayana Department of Biotechnology and Medical Engineering, National Institute of Technology, Rourkela Odisha, Pin: 769008, India Email: [email protected] Phone: +917750826049 Mr.Senthilguru Kulanthaivel Department of Biotechnology and Medical Engineering, National Institute of Technology, Rourkela Odisha, Pin: 769008, India Email: [email protected] Phone: +917735671699 Mr.Uvanesh Kasiviswanathan Department of Biotechnology and Medical Engineering, National Institute of Technology, Rourkela Odisha, Pin: 769008, India Email: [email protected] 1 ACS Paragon Plus Environment


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Dr. Kunal Pal Department of Biotechnology and Medical Engineering, National Institute of Technology, Rourkela Odisha, Pin: 769008, India Email: [email protected] Phone: 0661-2462289 Dr. Supratim Giri Department of Chemistry, National Institute of Technology, Rourkela Odisha, Pin: 769008, India Email: [email protected] Phone: +919438501472 Dr. Tapas K. Maiti Department of Biotechnology, Indian Institute of Technology Kharagpur West Bengal, Pin: 721302, India Email: tkmaiti@ hijli.iitkgp.ernet.in Phone: +919474597751 *Author for correspondence: Dr. Indranil Banerjee Department of Biotechnology and Medical Engineering, National Institute of Technology, Rourkela Odisha, Pin: 769008, India E-mail: [email protected] Phone: +919438507035

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Abstract

Success of bone tissue engineering (BTE) relies on the osteogenic micro-architecture of the biopolymeric scaffold and appropriate spatio-temporal distribution of therapeutic molecules (growth factors, drugs etc.) inside it. However, the existing technologies have failed to address both the issues together. Keeping this perspective in mind, we have developed a novel 3D implant prototype by stacking hexagonal close packed (HCP) layers of calcium alginate beads. The HCP arrangement of the beads lead to a patterned array of interconnected tetrahedral and octahedral pores of average diameter of 142.9 and 262.9 µm respectively, inside the implant. The swelling pattern of the implants changed from isotropic to anisotropic in z-direction in the absence of bivalent calcium ions (Ca+2) in the swelling buffer. Incubation of the implant in simulated body fluid (SBF) resulted in 2.7 fold increase in the compressive modulus. The variation in the relaxation times as derived from the Weichert's viscoelasticity model predicted a gradual increase in the interactions among the alginate molecules in the matrix. We demonstrated the tunability of the spatio-temporal drug release from the implant in a tissue mimicking porous semi-solid matrix as well as in conventional drug release set up by changing the spatial coordinates of the 'drug loaded depot layer' inside the implant. The therapeutic potential of the implant was confirmed against E. coli using metronidazole as the model drug. Detailed analysis of cell viability, cell cycle progression and cytoskeletal reorganization using osteoblast cells (MG-63) proved the osteoconductive nature of the implant. Expression of differentiation markers like alkaline phosphatase, runx2 and collagen type1 in human mesenchymal stem cell in vitro confirmed the osteogenic nature of the implant. When tested in vivo, VEGF loaded implant was found capable of inducing angiogenesis in a mice model. In conclusion, the bead based implant may find its utility in non-load bearing BTE.

Keywords Tissue engineering; Calcium alginate; Beads; Implant; Micro-patterned; Bone; Angiogenesis; Drug delivery.



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1. Introduction

In recent years, bioactive bone implants are gaining acceptance as a potential therapeutic alternative of autologus and allogenic bone grafts

1-2

. These implants when placed in vivo,

interact with the surrounding tissue environment and assists in restoring the functionality of the damaged bone. For the load bearing bones, bioactive implants are generally made up of metal and ceramics with a surface modification or coating of bioactive molecules such as cell adhesion proteins, peptides, growth factors and drugs

3-4

. On the other hand, for non-load bearing (e.g.

maxillofacial or craniofacial) or trabecular bone tissue engineering, therapeutics loaded biopolymeric scaffolds are used 5. The success of these 'bio-polymeric scaffold' assisted bone tissue engineering in non-load bearing bones greatly relies upon their micro-architecture and spatiotemporal distribution of therapeutic molecules at the site of injury. Various studies have confirmed that the pore size, pore distribution and porosity of the scaffold govern the osteogenesis in vivo

6-7

. Moreover, desorption kinetics of bone by osteoclasts is also dependent

on the aforesaid scaffold parameters. Another set of pertinent studies have independently shown that the abundance and the distribution of different growth factors (e.g. bone morphogenic proteins (BMPs), vascular endothelial growth factor (VEGF), and basic fibroblast growth factor (bFGF)) also control the final outcome of bone repair and healing, both in vitro and in vivo 8-9. With the advancements in the understanding of bone healing process, the researchers have emphasized on the definite micro-architecture of the scaffold and on the controlled spatiotemporal delivery of therapeutics at the injury site for efficient healing 3, 8. A feasible approach to achieve both the goals is to develop a scaffold fabrication technique that could ensure a tunable and reproducible patterned micro-architecture. To meet the aforesaid technical challenges, initial attempts were made to develop micro-patterned scaffolds using conventional high-end technologies such as computed tomography-guided fused deposition modeling, electrospinning, 3D printing, photolithography, rapid prototyping etc

4, 10-11

. Recently, more sophisticated hybrid

technologies have been explored for developing scaffolds with a defined micro-architecture and chemical environment. Skylar-Scott et. al. has developed P-selectin patterned 3D microfabricated scaffold using multi-photon photolithography

12

. A micro-patterned scaffold, with

open micro-channels of definite diameter was prepared by Varoni et. al. through electrochemical replica deposition of chitosan

13

. In 2015, Yuan et. al. reported the fabrication of ultra-thin 4

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scaffold by stable jet-electrospinning

14

. However, these techniques are very expensive and

require highly skilled professionals. Moreover, these fabrication processes are often found detrimental to the therapeutic molecules. In this regard, another promising but less explored approach is the use of polymeric beads for construction of 3D scaffold with a definite microarchitecture. This approach relies on either creation of micro-pores of definite size by using resolvable microspheres as porogen or by assembling beads in 3D fashion

15-16

. The use of

resorbable microspheres although ensures the micro-pores of definite size (analogues to salt leaching technique) but their spatial arrangement inside the scaffold is difficult to control. On the other hand, the potential advantages of the bead assembled system is that it enables the researchers to create definite microenvironment by simple manual maneuvering of the beads without the use of any sophisticated tools

17

. These beads could also act as a depot for a

controlled spatio-temporal distribution of therapeutics within the scaffold and surrounding injury site. To the best of our knowledge, till date, no comprehensive study has been carried out to explore both these possibilities in a single system. Keeping this perspective in mind, herein, we have reported the design, fabrication and characterization of a novel patterned 3D implant composed of alginate beads. Alginate is a well accepted biopolymer and has extensively been used for enzyme immobilization, cell encapsulation, drug delivery and tissue engineering applications 18. Chemically, alginate is water soluble, linear, poly-ionic polysaccharide consisting of alternating blocks of β (1→4) linked dmannuronic acid and α (1→4) linked l-guluronic acid residues 18. In our earlier studies, we have noticed that the calcium alginate beads of uniform size when placed in a water droplet, rearranged themselves into hexagonal close packed (HCP) structure according to the curvature of the droplet (data unpublished), which may be due to the surface tension of the system. Thus, here, we have explored that particular property of the water droplet-alginate bead physical system for the preparation of patterned 3D implant. For the patterning, two dimensional layer templates were first prepared by arranging the alginates beads in HCP and subsequently stacking these layers to achieve desirable three dimensional features. HCP arrangement is one of the well studied packing arrangements in crystallization, molecular self assembly and colloids. However, so far, no reports are available on the organization of alginate macrosphere into HCP and its subsequent application in the design of micro-patterned implants. These implants were further subjected for physico-chemical (SEM, swelling, mechanical and drug release studies) and 5 ACS Paragon Plus Environment


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biological (antibacterial, bone cell compatibility, cell cycle analysis, cytoskeletal organization and VEGF secretion) characterizations. Osteogenic properties of the implant was tested in vitro using human mesenchymal stem cell. Finally, in vivo performance of the implant was evaluated in mice model.

2. Materials and methods

2.1 Materials Sodium alginate (molecular weight: 7.72 x 104 g/mol, degree of polymerization: 476, M/G ratio 1.08) was bought from SDFCL, Mumbai, India. Calcium chloride (fused) was purchased from MERCK, Mumbai, India. Glutaraldehyde (25% aqueous solution) was procured from LOBA Chemie, Mumbai, India. Dulbecco’s Modified Eagle Media (DMEM), Dulbecco’s phosphate buffer saline (DPBS), Trypsin EDTA solution, Fetal Bovine Serum (FBS), antibiotic-antimycotic solution, metronidazole, MTT assay kit and nutrient agar were purchased from Himedia, Mumbai, India. RNAse, propidium iodide (PI), TRITC-Phalloidin and DAPI were procured from Sigma-Aldrich, India. VEGF was bought from Peprotech, India. Glycerol stock of E. coli was kindly provided by Prof. K. Pal, NIT Rourkela. MG-63, a human osteosarcoma cell line, was procured from NCCS, Pune, India. Bone marrow derived human mesenchymal stem cells were procured from Invitrogen, India.

2.2 Methods

2.2.1. Preparation and characterization of calcium alginate beads Calcium alginate (CA) beads were prepared by ionic gelation method as described in Agarwal et al 18. In brief, 1.5 % sodium alginate was dropped into crosslinking solution containing calcium chloride (2 % w/v) and glutaraldehyde (0.25 % v/v) using a 30G syringe. Preparation of rhodamine loaded bead was done by adding 100 µl of rhodamine stock solution (10 mg/ml) into 10 ml of alginate solution followed by ionotropic gelation. The beads were visually inspected and their average size was calculated by image analysis using fluorescent images. The structure of the dry bead was further observed under FESEM (Nova NanoSEM450) at 3kV. Prior to microscopy, beads were sputter-coated (Quorum Technologies, Q150RES) with gold. The 6 ACS Paragon Plus Environment

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calcium content of the beads was measured by EDAX. Swelling of the beads was studied in PBS for 24 h. For swelling analysis, 10 number of beads were used. The extent of swelling was calculated from the equation 1 given below.

Percentage swelling =

Wt − W0 ×100 W0

(1)

where, Wt is the weight of the samples after 't' h of swelling and W0 is the weight of the sample at '0' h. Biodegradation of the alginate beads was tested in vitro following the protocol reported by Boontheekul et. al. with little modification 19. Here, instead of using fresh media, we used MG63 spent media to provide biologically active degradation environment. In brief, prior to the experiment, beads were dried and weighted. Beads were then incubated in MG-63 spent media (72 hour old) at 37° C for 14 days. At definite time interval beads were taken out (10 beads at a time), dried and weighted again. The percentage of biodegradation was calculated using equation 2. Percentage biodegradation =

Wtd − W0 d x100 W0 d

(2)

where, Wtd is the dry weight of the beads after 't' h of incubation in degradation media and W0d is the dry weight of the beads at '0' h.

2.2.2. Preparation of the implant The implants were fabricated by placing the CA beads in a layer-by-layer arrangement (Figure 1). Beads were arranged in HCP configuration over the glass slide to form a layer of desired dimension (here, dimension of the base layer was kept 1.0 x 1.0 cm2 in most of the cases). Over it, 200 µl of sodium alginate solution (1.0 % w/v) was poured and crosslinked by 200 µl of calcium chloride solution (2 % w/v). Then, another layer of beads was placed over it and the process was repeated. The structure, thus, formed was dipped in 1 % sodium alginate and then immersed in crosslinking solution (containing 2 % calcium chloride and 0.25 % glutaraldehyde solution) for another 30 min. Thereafter, the implant was treated with glutaraldehyde neutralizing solution containing 0.1 M glycine and 1 mM calcium chloride for another 2 h. Following the above mentioned procedure, three and five layered implants were prepared and used in the studies. The morphological characteristics of the dried implant were examined using field 7 ACS Paragon Plus Environment


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emission scanning electron microscope (FESEM) (Nova NanoSEM450) at 5 kV post gold sputter coating (Quorum Technologies, Q150RES).The patterned micro-architecture of the implant was investigated using a stereomicroscope (Optika).

Figure 1 Schematic representation of the fabrication of the bead based implant.

2.2.3. Swelling and biodegradation analysis The swelling analysis of dry and wet implants was carried out in phosphate buffer saline (PBS, pH 7.4) and in incomplete DMEM media at 37 oC. For this, the implants were weighed and immersed in 25 ml of the solution separately. At definite time, the implants were carefully removed and weighed. The percentage swelling was measured using the equation 1 given above.


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Moreover, swelling dependent variations in the dimensions of the wet implant were also measured using a digital vernier caliper (Fisher Scientific, Accuracy: ± 0.03 mm/0.001 inches). The measurements were taken at least at 3 random positions for each dimension and the average values were reported. Biodegradation of the implant was tested in vitro using spent media of osteoblast cells following the same protocol adopted for alginate beads (refer section 2.2.1).The percentage of biodegradation after 14 days of incubation was calculated using equation 2. In this case, Wtd was the dry weight of the implant after 14 days of incubation in degradation media and W0d is the dry weight of the implant at '0' h. In addition to checking the final weight loss, time dependent variation in the dimension of wet implant was also measured using the digital vernier caliper following the protocol mentioned above.

2.2.4. Mechanical characterizations The mechanical properties of the implants were measured in compression-stress relaxation mode using a static mechanical tester (TA.HD Texture analyzer, Stable Micro Systems). The study was conducted by vertically compressing the samples to 10 % strain (without breakage) against a trigger force of 3 g-force using a flat probe (30 mm diameter and crosshead speed 1 mm/s) and further holding the probe for 60 s. For this, cubical samples of dimension 20 (width) x 20 (length) x 10 (height) mm3 were used. To understand the time dependent variation in the viscoelastic behavior of the implant, all the samples were kept in SBF and the analysis was carried out at three definite time points (Day 1, 7 and 14). The percentage stress relaxation (% SR) was calculated by using equation 3 given below.

%SR =

F0 − FR ×100 F0

(3)

where, F0 is the maximum force attained during the compression stage and FR is the residual force at the end of the relaxation period. The compressive modulus of the implant 'EC' was calculated by equation 4 given below using the data point taken from the linear region of the graph during compression stage.

EC =

σ max ∆lz

(4)

where σmax is the maximum stress and ∆lz is the change in length along z axis.



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The viscoelastic parameters of the implants were further predicted by fitting the stress relaxation data in the Weichert model. In this study, the model was created using 3 dashpots, which provides three relaxation times. The mathematical equation for this model has been given below.

P(t ) = P0 + P1.e−t/τ1 + P2.e−t/τ2 + P3.e−t/τ3

(5)

where, P0, P1, P2,P3 are the spring constant; τ1, τ2, τ3 are the time constants of the dashpots and 't' was the time.

2.2.5. Drug release analysis In order to characterize the pattern of the drug release from the implant in tissue mimicking semi-solid porous matrix, rhodamine loaded implants were prepared. For the analysis, both three and five layered implants were used. In both the cases, a single HCP unit of rhodamine loaded beads (2D, number of beads = 7) was placed at the center of the middle layer of the implant. Thereafter, the implants were placed at the center of a petri-plates containing solidified 0.6 % agarose gel and incubated at 37 oC in a closed humidified chamber. Time dependent release of rhodamine and its subsequent diffusion through the agarose gel was analyzed by taking images using camera (Canon A2400 IS) in a gel documentation system and subsequently processing them with MBF ImageJ software. The release of the drugs from the implant in solution was analyzed following the method previously described by Pakzad et. al.

20

. For this purpose, metronidazole loaded bead was

prepared first following the same protocol adopted for the preparation of rhodamine loaded beads (section 2.2.1). Initial concentration of metronidazole used for this purpose was 2mg/ml. The drug loaded implant was prepared by

placing single hexagonal close pack unit of

metronidazole loaded beads at the centre of the implant. In brief, metronidazole loaded implants were immersed in 50 ml PBS (pH 7.4). At definite time intervals, 3 ml of the supernatant was withdrawn from each set and corresponding drug concentration was measured using UV–visible spectrophotometry. Herein, metronidazole (λmax = 321 nm) was taken as a model drug. The study was performed in triplicates using three and five layered implants. The loading efficiency was analyzed by crushing dried implants and resuspending the powder in 10 ml of PBS (pH 7.4) for 24 h. Thereafter, the drug concentration in the supernatant was measured using spectrophotometry. The release data was further fitted into Korsmeyer Peppas model

21-22

.

Equation 6 was used for the modeling of the released data. 10 ACS Paragon Plus Environment

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Mt = k.t n M∞

(6)

where, ' Mt/M∞ ' is the fractional drug release at time 't', 'k' is the release rate constant and n is the release exponent. Furthermore, in vitro assessment of the therapeutic delivery potential of the implants was analyzed by antibiotic susceptibility test as described elsewhere

23

. In brief, 100 µl of E. coli

culture (2 × 106 CFU/ml) was spreaded over nutrient agar plates and metronidazole loaded sterile implants were placed in the center of the plates. The plates were incubated overnight at 37 °C and corresponding zone of inhibition (ZOI) was measured thereafter.

2.2.6. Biological characterization of the implant The biological performance of the implants was evaluated against MG-63 cells. MG-63 cells were maintained in DMEM with 10 % heat inactivated FBS in a humidified (95 %), CO2 incubator (5 %) at 37 oC with regular passages at 80-90 % confluency. For all the biological characterizations, implants were fabricated under sterile conditions [refer section 2.2.2] and equilibrated with DMEM media. The cells were seeded into the implant at a concentration of 1 x 106 cells/ml. A preliminary evaluation of the viability and time dependent in-growth of cells inside the implant was done using MTT assay with little modification. Here, we incubated the cell loaded implant with MTT reagent for the formation of formazone crystal but at the end we did not add DMSO (a reagent responsible for solublilization of MTT . This was done to characterize the 3D distribution of purple colour formazone crystal inside the implant which actually corresponds to cellular location. Flow cytometry based live-dead assay and cell cycle analysis were performed after 72 h of initial cell seeding using a table top flow cytometer (Accuri C6, BD Biosciences). Tissue culture plate (TCP) and calcium alginate hydrogel were taken as controls. Variations in the cellular morphology were examined using fluorescence microscopy (Olympus). For this purpose, cells were fixed with 4 % paraformaldehyde and stained with TRITC-Phalloidin and DAPI. Furthermore, alkaline phosphatase (ALP) and vascular endothelial growth factor (VEGF) secretion by MG-63 cells was estimated using ALP assay kit (Span Diagnostics) and VEGF ELISA kit (Abcam) respectively following manufacturer’s instructions.



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Osteogenic potential of the implant was tested using human bone marrow mesenchymal stem cell (hMSC). In brief, hMSCs were maintained in DMEM low glucose supplemented with 10 % FBS and 10 ng/ml bFGF in a humidified (95 %), CO2 incubator (5 %) at 37 oC. hMSC were then trypsinized and seeded on to the implant at a concentration of 1x106 cells/ml and cultured for 14 days in osteogenic differentiation media (DMEM high glucose supplemented with 10 % FBS, 1X GlutaMAX, 1X NEAA, 10 mM beta-glycerophosphate, 10 nM dexamethasone, 50 µM ascorbic acid and 10 ng/ml BMP2) with a change at regular interval of two days. TCP and pure alginate gels were taken as control. At different time interval, 10 µl of the supernatant was collected for ALP expression analysis. At day 14, cells were trypsinized, collected through centrifugation at 1000g and one part of the cells was lysed and subjected for RT-PCR using an one step RT-PCR kit (Qiagen) for analysis of osteogenic differentiation markers namely runx2 and collagen type I. The primers used for the analysis were Runx2 forward 5’ TTTACTTACACCCCGCCAGTC 3’, reverse

5’

CAGCGTCAACACCATCATTCTG

3’;

collagen

type

I

forward

5’

AGACTGGCAACCTCAAGAAGGC 3’, reverse 5’ CGGGAGGTCTTGGTGGTTTTGT 3’ and GAPDH

forward

5’

CATGAGAAGTATGACAACAGCCT

3’,

reverse

5’

AGTCCTTCCACGATACCAAAGT 3’. Remaining cells was evaluated for re-plating efficiency and subsequent osteogenic commitment. For this, concentration of live cells was determined by trypan blue assay using a hemocytometer. Then, the cells were seeded on a new 24 well tissue culture plate and allowed to adhere for 24 h. After 24 h, non adherent cells were taken out from each well and counted using hemocytometer. Plating efficiency was measured as percentage of cells adhered with respect to total cell seeded. Morphology of the replated cells was further investigated using fluorescence microscopy (Olympus). For this purpose, cells were fixed with 4 % paraformaldehyde and stained with FITC-Phalloidin and DAPI. Osteogenic commitment of the replated cells was evaluated by alizarin red S staining as per standard protocol.

2.2.7. In vivo assessment of the performance of the growth factor loaded implant Performance of the VEGF loaded implant was tested in vivo in a mice model following a protocol reported elsewhere

24

. For this purpose, an implant containing 500 ng VEGF loaded

alginate bead (without glutaraldehyde crosslinking) positioned at the centre of the implant was used. 3 healthy Swiss albino mice (4 -6 week old) of average body weight 22-24 g were selected for this study. VEGF loaded cylindrical wet implant of average diameter 6.5 mm and height 2.5 12 ACS Paragon Plus Environment

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mm was implanted in the dorsal subcutaneous pocket (right side) through a small surgical incision. Similar kind of implant without growth factor was inserted in the dorsal subcutaneous pocket (left side) of the mice following the same procedure. All the operations were done with the prior approval of the Institutional Animal Ethical Committee of IIT Kharagpur (IEAC file number - IE-5/TKM-BT/3-16). During the operation mice were anesthetized using a standard dose of ketamine-xylazine. Post operation, individual mouse was housed separately in aseptic environment with food and water ad libitum. After 7 days of operation, mice were sacrificed through cervical dislocation and the inner layer of the skin that was in contact to the implant was examined. The zone of implant contact was visually investigated to find out extent of angiogenesis at the region of interest and imaged with digital camera (ASUS, 13 MP).

2.2.8. Statistical analysis All the experiments were carried out in triplicate and the data were expressed as Mean ± S.D. Statistical significance of the data was evaluated using single variance ANOVA under 95 % confidence interval.

3. Results and discussion

3.1. Preparation and characterization of CA beads CA bead has already been well established as a cell encapsulation and therapeutic delivery vehicle

18, 25-27

. A number of research groups have reported the preparation of CA beads of

varying dimensions following different techniques such as syringe extrusion emulsification coupled with internal gelation

28-29

and microfluidics

18

, membrane

30-31

. All these techniques

involve ionic gelation of the alginate molecules in presence of bivalent/trivalent ions such as Ca+2, Mg+2 or Al+3 ions. In the present study, we employed syringe extrusion methodology to prepare the calcium alginate beads. The average size of the swollen beads was 1.27 ± 0.07 mm (Figure 2 A, Inset) with a percentage yield of 96.13 ± 1.30 %. Preliminary physical inspection ensured that the beads were spherical in shape. SEM analysis further showed that the dry beads had rough surface topology (Figure 2 A). The elemental analysis by EDAX confirmed significantly higher content of calcium (11.09 ± 1.03 %) in CA beads than that of sodium alginate (1.29 ± 0.41 %), suggesting Ca+2 mediated ionic gelation of the alginate matrix (Figure 2 13 ACS Paragon Plus Environment


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B). The calcium content is a crucial parameter in the context of mechanical strength of the beads. Analysis of the swelling of the beads showed a small yet significant change over a period of 24 h (p < 0.05). Percentage swelling of the beads at the end of 12 h was 15.1 ± 2.1 %, which reduced to 13.6 ± 4.2 % in the next 12 h of analysis (Figure 2 C). Such a reduction in swelling may be due to the leaching of the bound Ca+2 ions, resulting in the erosion of alginate matrix. Biodegradation of alginate based scaffold or implants is a serious concern for its biomedical application. Absence of the enzyme 'alginase' in mammalian system prevents degradation of alginate polymer chain in vivo. However, ionically crosslinked alginate chains are often prone to dissolution both in vitro and in vivo through the replacement of the bi- or multi-valent ions responsible for ionic gelation by mono-valent ions or through chelation

32

. This leads to mass

loss of alginate structure through 'erosion'. Our result showed that the extent of biodegradation of alginate beads over two weeks was approximately 20 % (Figure 2 D). The percent degradation of beads was 14.30 ± 0.98 and 19.82 ± 1.30 % at day 7 and 14 respectively.


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Figure 2 Characterization of the CA beads. (A) Scanning electron micrograph of the dried CA bead. Inset: fluoroscence image of the rhodamine loaded beads arranged in 2D hexagonal close packed format (B) Analysis of the calcium content (w %) in CA beads by EDAX. Sodium alginate (SA) was taken as reference. (C) Swelling analysis of CA beads in phosphate buffer saline (pH 7.4) at 37 oC. The study was carried out for 24 h. (D) Biodegradation calcium alginate beads in cell culture media. All the studies were performed in triplicate and data was expressed as Mean ± S.D.

3.2. Fabrication of the implant Fabrication of a bead based implant with a definite microarchitecture requires 3D spatial patterning of the beads. Herein, we first patterned the beads in HCP arrangement to form a 2D layer of dimension 1 x 1 cm2 taking the advantages of surface tension driven reorganization of beads in water droplet. Later, we stacked similar layers one over the other to achieve a desirable 3D configuration and glued it with small amount of sodium alginate solution which made a thin binding layer around the stacked beads (Figure 3 A1-3). This packing was important to maintain the overall structural integrity of the implant. The aforesaid procedure allowed us to tune the dimensions of the implant as per the requirement by modulating either (i) the dimensions of the beads, or (ii) the dimensions (length and breadth) of each bead layer, or (iii) the number of stacking (height). Here, we prepared 3-layered and 5-layered (designated as 3L and 5L) implants of average dimensions of 1.11 ± 0.07 x 1.05 ± 0.08 x 0.81 ± 0.08 and 1.15 ± 0.08 x 1.13 ± 0.06 x 1.05 ± 0.10 cm3, respectively. The fabricated structures were stable, consolidate and easy to handle. Upon air drying at room temperature, the implant shrank into a small cubical structure of dimensions 5.32 ± 0.25 x 5.11 ± 0.19 x 2.67 ± 0.16 mm3 with a very compact integrity. The scanning electron microscopy of dried implant also demonstrated the intact HCP arrangement of the beads (Figure 3 B1). The average bead diameter in the dried implant was 696.51 ± 60.72 µm. The dried implant appeared solid without any pores on its surface. At the center of the implant, the beads retained their smooth shape; while, at the edges, beads had wrinkled morphology. Some cracks were observed on the implant surface, which might have developed during sample processing for SEM (Figure 3 B1, Inset). Once swelled in PBS, the structure was able to regain



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its original shape with an intact bead arrangement. Such property implies that the implant could also be stored in dry form before use. In HCP, the spherical units repeat themselves after every two layers, creating "a-b-a-b-a" kind of structural arrangement ('a' and 'b' stands for two adjacent layers). Such arrangement of the spheres would generate two different kinds of pores, namely, octahedral and tetrahedral pores 3334

. It has already been established that if spheres of radius ‘R’ is arranged in HCP, then the radius

of a hypothetical sphere that could just fit into the tetrahedral pore will be 0.225R and the same for octahedral pore will be 0.414R 34. Moreover, the pores in HCP follow a repetitive pattern and remain connected throughout the structure. As per this consideration, our implants would have interconnected repetitive pores of average diameter 142.9 ± 7.5 and 262.9 ± 13.9 µm. Such interconnectivity of the pores is essential for efficient transport of nutrients and other soluble factors. Furthermore, HCP ensures ~26 % void volume (maximum) which can be correlated with the porosity of the implant. Analysis of the bead implant by stereomicroscope showed an arrangement of the micropores corresponding to the HCP arrangement of the beads (Figure 3 B2). It was evident from the figure that all the pores were interconnected (dotted red line in figure 3 B2). Depending upon the orientation of the beads in the adjacent layers, tetrahedral or octahedral pores would be generated from the representative 2D template shown in Figure 3 B2. Figure 3 B3 and B4 showed the schematics of the octahedral and tetrahedral pores. Many reports have already emphasized on the use of macroporous scaffolds with interconnected heterogeneous pores of size ~200-900 µm for bone tissue engineering

35-36

. A comparison of our implant with

the above mentioned scaffolding criteria for bone tissue engineering clearly suggests its microstructural parity. Notably, due to the intrinsic robustness and consistency of HCP organization, the bead based scaffolding technique is associated with high reproducibility (only consistency in bead size needs to be maintained). It also allows researcher to control the pore size by simply controlling the bead diameter, as the later is directly proportional to the bead diameter.


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Figure 3 Fabrication of the implant. (A1) Arrangement of CA beads in layers with rhodamine loaded beads at the center, (A2) top and (A3) side view of the implant encased in the thin layer of alginate. Scale bar represents 10 mm. (B1) Scanning electron micrograph of dried implant. The green dashed lines demonstrates intact hexagonal close packing of the beads in dried implant. Inset: Scanning electron micrograph of the inter bead association in the dried implant. (B2) Study of the arrangement of the micropores inside the implant by stereomicroscope. (B3) and (B4) showed the schematics of the octahedral and tetrahedral pores.

3.3. Swelling and biodegradation analysis Swelling behavior is an important criterion for evaluating the functional characteristics and the applicability of a bio-polymeric implant. The swelling involves progressive interaction of the polymer matrix with solvent molecules until equilibrium is attained. We performed the swelling study for both the dry and wet implants. To understand the role of Ca+2 ions present in the swelling media, we used PBS and incomplete DMEM media (5 mM Ca+2) for the study. Study showed that the extent of swelling of both dry and wet implants was higher in DMEM than in PBS (Figure 4 A1-2). The swelling of the dry implant in DMEM was 580.25 ± 116.05 %, while the same in PBS was 442.7 ± 61.93 % after 24 h. Thereafter, a decrease in swelling in PBS was



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observed in case of dry implant over time. This could probably be due to the release of Ca+2 from the matrix thereby resulting in the degradation of the matrix structure. In the wet implant, the percentage swelling in PBS and in DMEM after 48 h was 8.97 ± 2.28 and 27.3 ± 3.0 %, respectively. A critical analysis revealed that in the wet implants, variation in swelling in DMEM and PBS was statistically significant for all the data points (p < 0.05), while in case of the dry implant, a similar observation was also noted for most of the data points. The wet implants showed a decrease in the swelling in PBS during the initial 3 h of analysis, which might be due to the dissolution of the loosely bound alginate polymer from the implant. The higher swelling of the implant in DMEM may be due to absorption of the different charged molecules such as amino acids on the polymer matrix, owing to electrostatic repulsion among the alginate chains. Furthermore, analysis of the relative changes in the implant dimensions during swelling revealed that the implants followed an anisotropic swelling behavior in PBS (Figure 4 B1-2). The relative variation in the length, breadth and height of the implant after 48 h of swelling analysis were in ratio 0.98 : 0.99 : 1.06. The variation of dimension along z-axis to that of x- or y-axis was found statistically significant (p < 0.05). However, in DMEM, implants showed an isotropic swelling behavior with dimensional ratio of 1.21 : 1.19 : 1.22. Such anisotropic swelling of the implant structure in PBS may be due to the low calcium concentration. This was further supported by the complete disruption of the implant after 90 h of incubation in PBS (pH 7.4). Biodegradation of the implant was checked for 14 days in the spent media of MG-63 cells. During this time period, 26.79 ± 2.5 % decrease in the volume along with 12.154 ± 0.68 % loss in mass of the implant was observed. As mentioned earlier (section 3.1), it could be believed that such degradation happened due to erosion of the polymer molecules.


19


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firmness of the system, was lowest (41.4 ± 4.4 g-force) at the beginning (day 1), reached maximum at day 7 (110.1 ± 11.8 g-force) and then decrease to 95.4 ± 13.4 g-force at day 14. Time dependent variation in residual force (FR) also followed similar pattern. The % SR was found significantly higher at day 1 (69.0 %) which reduced to ~63 % over time (day 7 and 14) (Table 1). The result clearly implied that the rigidity of the implants increased over time which could probably be due to gradual ionotropic gelation of some free alginate molecules present in the core. The constituent bivalent cations of SBF (e.g. Mg+2, Ca+2) probably diffused into the implant over time and crosslinked the free alginate molecules. Such crosslinking ensured enhanced polymer-polymer interactions and resulted in a decrease in the stress relaxation at day 7 and 14 in comparison to day 1. In addition, incubation in SBF also causes mineralization of the scaffolds which may increase its rigidity and compressive modulus 39. A small decrease in the F0 and FR values from day 7 to day 14 could be explained in terms of reversible acetal crosslinking of glutaraldehyde 40. It may happen that number of acetal bonds formed initially got cleaved over time making the implant structure weak. Compressive modulus of the implants was 0.17 ± 0.01, 0.46 ± 0.05 and 0.40 ± 0.06 kPa at day 1, 7 and 14 respectively (Table 1). These values were less than that of average compressive modulus of the cancellous bone (2-20 MPa) 3. However, literature suggests that a number of hydrogels with low compressive modulus have been used in bone tissue engineering with success especially in case of non-load bearing bones 41-42. Modeling of normalized relaxation profiles using the Weichert's 3 dashpot-element mechanical model of viscoelasticity revealed only 1.16 and 1.06 fold increase in the instantaneous relaxation time (τ1) at day 7 and 14 with respect to day 0 (Table 2). Instantaneous relaxation time generally corresponds to molecular rearrangement in the material under stress. In our case, uniformity in these values signifies the absence of any molecular rearrangement in the bead material over time. On the other hand, the intermediate and the delayed relaxation times (τ2 and τ3) provide information about the breakage of polymer–polymer interaction and polymer chains respectively. Herein, about two fold increase in both values was observed at day 7 and 14 in comparison to day 1. This supports our arguments in favor of the prolonged ionotropic gelation of the uncrosslinked alginate molecules in SBF. An increase in the mechanical strength of the implant over time when kept in simulated physiological fluid clearly implied that it could effectively be used in vivo.


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Figure 5 (A) Stress relaxation profile of implants. The implants were incubated in SBF for different time interval prior to the experiment. Black, red and blue lines represent relaxation profiles of implant after 1, 7 and 14 day of incubation respectively. Weichert’s model fitting of the relaxation profile of the implant at day 1 (B1), day 7 (B2) and day 14 (B3).

Table 1 Time dependent variation in the mechanical properties of the implant Compressive modulus (EC)

F0 (g-force)

FR (g-force)

% SR

Day 1

41.4±4.4

12.9±3.1

69.0±3.2

0.17 ± 0.01

Day 7

110.1±11.8

41.15±5.3

62.6±0.8

0.46 ± 0.05

Day 14

95.4±13.4

30.1±7.1

63.3±2.3

0.4 ± 0.06

(kPa)



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Table 2 Values of instantaneous, intermediate and delayed parameters obtained from the Weichert model. P0, P1, P2 and P3 are the spring constants (normalized value), τ1, τ2 and τ3 are the time constant of the dashpots. R2 is the coefficient of regression.

P0

P1

τ1(s)

P2

τ2(s)

P3

τ3(s)

R2

Day 1

0.27

0.25

31.8

0.22

0.47

0.25

3.17

0.99

Day 7

0.43

0.3

36.9

0.08

1.06

0.18

6.07

0.99

Day 14

0.43

0.3

33.8

0.11

1.06

0.18

5.87

0.99

3.5. Drug release study We analyzed the spatio-temporal release of the therapeutic molecules from the implant in a tissue extracellular matrix (ECM) mimicking environment taking rhodamine as a model drug. For this purpose, the drug loaded implant was placed on 0.6% agarose gel and the drug transport from the implant to the gel was monitored by fluorescence imaging (Figure 6 A1-2). 0.6% agarose gel has often been used as a model tissue matrix for its mechanical resemblance (strength and elasticity) with native tissues

43

. Analysis showed that rhodamine from the source (beads present at the

central position of the middle layer), first diffused inside the implants (initial 24 h) resulting in an overall increase in the fluorescence intensity at the center of the top layer (P0) in case of both, 3L (from 150.38 to 162.04 a.u.) and 5L (from 137.85 to 154.49 a.u.) implants. Within this time period, a portion of the loaded rhodamine also diffused to the surrounding agarose gel. An increase in fluorescence intensity (corresponding to rhodamine concentration) at distal position 'P35' was observed in both, 3L (34.91 to 45.63 a.u.) and 5L (34.17 to 42.73 a.u.). Over time, the zone of diffusion increased following a continuous gradient of rhodamine. A significant variation in the spatial distribution profile of rhodamine was observed in 3L and 5L throughout the analysis. Within a specific time scale, implant with lower number of bead layers released more drug to the surrounding. This could be due to the lesser interstitial space in 3L than 5L which led to low intra-implant diffusion of rhodamine. The velocity of the moving dye front inside the agarose gel for 3L was significantly higher (average velocity: 0.333 ± 0.039 mm/hr) in comparison to 5L (average velocity: 0.193 ± 0.023 mm/hr) (p < 0.05). This could possibly be due to the presence of higher concentration gradient between the source and the boundary layers in 3L than 5L as mentioned above. At the end of day 5, intensity of the dye at P35 was 66.28 and 54.43 a.u. for 3L and 5L, respectively. This proves that such drug loaded bead implant could 22 ACS Paragon Plus Environment

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effectively be applied for delivery of therapeutics to the neighboring tissue site in a sustained manner upon in vivo implantation. Moreover, the spatio-temporal distribution of the therapeutics from the implant can be tuned effectively by just manipulating the number of bead layers. The distribution profiles of the rhodamine showed that there was no accumulation of dye at the implant-gel interface, which ensures its effectiveness as a delivery system. We further confirmed the drug release properties of the implant using conventional procedure taking metronidazole as the model drug 20. Here also, we observed a significantly higher release of metronidazole from 3L than 5L implant (p < 0.05) (Figure 6 D1). After 24 h, the cumulative percentage drug release of metronidazole for 3L and 5L was 73.28 ± 4.48 and 62.79 ± 4.76 %, respectively. The drug release profile was further fitted to different release models (zero order model, first order model, Higuchi model, Hixson-Crowell model and Korsmeyer- Peppas model. The best model fitting for both, 3L and 5 L were Higuchi and Korsmeyer- Peppas model (R2 > 0.95, Supplementary data, Table TS1). As per the Korsmeyer- Peppas model, the value of the release exponent ‘n’ for the 3L implants was 0.49 and the same for 5 layers was 0.40). Earlier, it was reported that for a swellable polymeric controlled release system (swelling should be less than 25% of its original volume), ‘n’ value can be used to predict the release mechanism. For Fickian diffusion the value of 'n' was found 0.5, 0.45 and 0.43 for thin film, cylinder and spherical samples respectively; whereas for case II transport, 'n' value would be 1, 0.89 and 0.85 respectively for the samples of aforesaid geometry. Any value of ‘n’ between its Fickian diffusion limit and case II transport limit implied the existence of Anomalous non Fickian transport

22

. In the present case, although the drug release profile fits well with Korsmeyer-

Peppas model but it is not appropriate to use 'n' value for the prediction of drug release mechanism from the implant (Figure 6 D2). This is primarily because assumptions of Korsmeyer- Peppas model were probably not valid for this system. In case of the implant, the drug was first release from the bead in to the interstitial fluid present in the void space. The drug molecules present in the fluid would have then two options, first it may diffuse into the non-drug loaded neighboring alginate beads or it may move out by crossing the outer alginate layer of the implant. In such case, the pattern of release will be too complex to determine. However, a simplified explanation could be the consideration of the existence of diffusion and ‘swelling– dissolution–erosion’ mechanisms which are common for many alginate based systems

44

.

Furthermore, when metronidazole loaded implants were placed onto agar plates; a clear ZOI was 23 ACS Paragon Plus Environment


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observed against E. coli, indicating the diffusional release of metronidazole (Figure 6 D1, Inset). The 3L and 5L implants showed a significant difference in the ZOI diameter of 29.33 ± 0.76 and 26.91 ± 1.25 mm respectively (p < 0.05), following a similar trend as that of rhodamine release.

Figure 6 Time lapse imaging of the release of rhodamine from 3L (A1) and 5L (A2) implants. Intensity profiling of the rhodamine gradient across the alginate gel over time in 3L (B1) and 5L (B2) implants in a 90 mm petri dish. The fluorescent intensity could be correlated with the rhodamine concentration at that point. Black, red, blue and pink lines correspond to intensity profile at day 0, 1, 3 and 5 respectively. Heat map representation of the fold change in the fluorescent intensity at different spatial coordinates over time in 3L (C1) and 5L (C2) implant. P10, P15, P20 and P25 corresponds to distance (in mm) from the center of the implants. Analysis of the metronidazole release from the implant in PBS (D1). Formation of zone of inhibition around the metronidazole loaded 3L and 5L implants when tested against E. coli (Inset D1). Korsmeyer Peppas (best fit) model (D2) of metronidazole release. Black and red represent 3L and 5L implants respectively. KP model analysis was done for initial 60% of cumulative drug release.


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3.6. Biological characterization of implants Alginate based 3D formulations, both scaffolds and hydrogels, have been tested in bone tissue engineering applications for couple of years. Some of the recent reviews have nicely documented the current status of the alginate based bone tissue engineering

45-46

. A critical analysis of these

reviews revealed that although the alginate based systems support the viability, proliferation and differentiation of bone cells to a varied extent depending upon their composition and physical features but none of them truly meet all the important functional requirements at a time especially the controlled spatio-temporal release of the therapeutics and a defined osteoconductive micro-patterned interior. In this regard, we have already proved the credibility of our bead based implant (refer section 3.2 and 3.5). To evaluate the biological performance of the implant, we first performed a preliminary analysis of the viability and in-growth of the MG-63 cells inside the implant using a modified MTT method

47

. The analysis was based on the semi-

quantitative mapping of the spatial distribution of the purple formazan crystal (formed by MTT reaction) inside the implant which corresponds to the presence of metabolically active cells at that location. The time dependent distribution pattern of the insoluble MTT crystals revealed that the cells migrated downwards from the top of the implant after initial seeding (Figure 7 A1-3). To confirm the cellular in-growth in a 5L implant, each layer was individually investigated (Figure 7 B1-5). The color intensity profiling using a semi-quantitative image based analysis revealed that after 5 days of initial cell seeding, cell population at the middle and the bottom layer was almost 75% and 55 % respectively, to that of the top layer (data not shown). A combined effect of migration and proliferation probably lead the cells to populate the implant interior after 5 days of culture.



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Figure 7 (A1-A3) Spatial distribution of the purple formazan crystal inside the 5L implant at day 1(A1), 3(A2) and 5(A3). The scale bars represent 5 mm. Distribution profile of purple formazan crystal indicates an in-growth of cells inside the implant. (B1-5) Presence and distribution of MTT crystals on each bead layer.

We further investigated the viability, cell cycle progression, and cytoskeletal reorganization of MG-63 cells cultured on the implant. For an appropriate comparison, we took calcium alginate hydrogel and tissue culture plate as control. MG-63 (osteosarcoma) is a well tested cell line for bone tissue engineering research

48

. Flow cytometry based live-dead assay after 72 h of initial

seeding showed maximum cell viability in case of implant (87.76%) followed by TCP (81.91%) and alginate gel (69.04%) (Figure 8 A). Low viability in case of alginate hydrogel could probably be due to its poor stiffness which is essential for bone cell adhesion and growth. On the other hand, the highest viability in the implant may be attributed to its osteoconductive micropatterned interiors. When compared with recently reported bio-printed 3D alginate scaffold, the cell viability was found to be almost in the similar range 49. The cell cycle analysis showed that there was no halt in the normal cell cycle progression of MG-63 cells in any case. Major fraction of the cell population were in G0/G1 phase (~69.36-72.89 %), followed by S phase (~12.6718.14 %) and G2/M phase (~10.49-13.41%) (Figure 8 B). A little reduction in S-phase cell


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population was observed in case of implant (12.67 %) and hydrogel (13.94 %) in comparison to TCP (18.14 %). The distribution profile and the cell-cycle data together confirmed that the implant favored the bone cell migration and proliferation. To confirm the existence of healthy cells, the inner layer of the bead based implant was manually exposed; cells were stained with TRITC-Phalloidin and DAPI and investigated under fluorescence microscope. Analysis revealed that a number of cells were able to maintain their spreaded and elongated morphology in the implant while most of them were found rounded on the alginate hydrogel (Figure 8 C). Analysis of cellular area demonstrated that the average spreading area of the cells cultured on implant (668.1 ± 305 µm2) was close to that of TCP (876.77 ± 198.3 µm2); whereas, the same for alginate gel was quite less (330 ± 143.3 µm2) in comparison to both TCP and implant (Figure 8 D). Statistical analysis showed that the variation in cell spreading area was insignificant between TCP and implant but significant between TCP and alginate gel. To further evaluate the bio-functionality of the implant, vascular endothelial growth factor (VEGF) secretion was quantified. VEGF is a survival factor for endothelial cells and the most potent inducer of angiogenesis. Angiogenesis is one of the most important cellular activities required for bone regeneration and repair. During physiological reconstruction of bone, the cells present at the wound site (including bone cells) secrete VEGF to ensure early establishment of vasculature 50. The present analysis demonstrated a significant variation in VEGF expression depending upon the culture systems (p < 0.05). The cells cultured on the implant showed 1.23 and 1.50 folds increase in the VEGF expression with respect to alginate hydrogel and 2D monolayer culture, respectively (Figure 8 E). This could be due to the development of hypoxic conditions inside the implant and hydrogel, which induce higher VEGF expression via activation of HIF-1α pathway

51

. After 72 h of seeding, cells

cultured on different substrate did not show any significant variation in ALP expression (Figure 8 F). This is probably because MG-63 is a differentiated osteoblast cell line, therefore it is less likely for them to show any variation in the expression of early differentiation marker like ALP. Interestingly, cellular expression of ALP in case of implant was comparable to that of TCP which suggested that implant environment help in maintaining their differentiated state. All these results together implied that the implants could support bone cell viability and functionality to a better extent in comparison to normal alginate hydrogels.



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Figure 8 Biological characterization of the implant. (A) Live-dead assay of the MG-63 cells. Live cell population have been enclosed in green box, while dead ones in red. (B) Cell cycle profiling of MG-63 cells. (C) Fluorescent micrographs of the cells stained with TRITCPhalloidin and DAPI. Scale bar represents 50 µm. (D) Average cellular area of the cells. (E) Relative change in the cellular VEGF expression. TCP and calcium alginate hydrogel was taken as controls. (F) Analysis of alkaline phosphatase (ALP) expression. ALP activity was measured in the supernatant of the culture. ‘*’ designates statistical significance with p < 0.05.

Osteogenic potential of the implant was further tested in vitro using hMSC. It has already been proven that the scaffold micro-architecture governs the osteogenic differentiation of hMSC

52

.

Osteogenic differentiation of hMSC is associated with a time dependent expression of various differentiation markers such as alkaline phosphatase (ALP), Runt-related transcription factor 2 (Runx2), osteocalcin (OCN), osteopontin (OPN), bone sialoprotein (BSP) and collagen type I (Col1). Among the osteogenic markers, ALP and Runx2 are considered as early osteogenic marker; whereas expression of Col1 is associated with bone cell mediated matrix mineralization. In this study, we checked the expression of ALP, Runx2 and Col1 (Figure 9 A and B). At day 7, 28 ACS Paragon Plus Environment

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ALP activity in implant was found slightly greater more than hydrogel and TCP but the data was not statistically significant. However on day 14, a significant increase of ALP activity was observed for implant (~1.35 fold with respect to hydrogel). RT-PCR study further confirmed higher expression of both the differentiation markers (Runx2 and col1) in case of cells differentiated on implant. After 14 days of culturing, the partially differentiated hMSCs were harvested from the matrices (TCP, hydrogel and implant) through trypsinization and replated on a fresh tissue culture plate. Analysis showed >90 % of cell adherence after replating (data not shown). Figure 9 C showed the morphology of the replated partially differentiated hMSCs (14 day osteogenic differentiation) after 48 h. From the figure, it is evident that cells cultured on the implant did not lose its adherence property and restored their characteristic well spreaded morphology. Another set of replated cells were cultured on the tissue culture plate for next 3 days in osteogenic medium and subjected for alizarin red staining which is a proof of osteogenic matrix deposition. Our study showed that that cells isolated from matrix resulting in higher matrix deposition (Figure 9 D). All these data clearly suggest that the implant support the osteogenic differentiation of hMSC. Such osteogenic nature of the implant could be attributed to its unique microarchitecture (pore size, porosity and 3D microenvironment) or presence of hypoxic zone inside the implant which may favor differentiation of hMSC 53.



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Figure 9 Study of the osteogenic properties of the implant in vitro. The study was carried out using human mesenchymal stem cells (hMSC). (A) Time dependent variation of alkaline phosphatase expression. ALP activity was measured in the supernatant of the culture. (B) Study of the expression of osteogenic marker through RT-PCR. The study was carried out after culturing the hMSC on different substrates in presence of osteogenic media for 14 days. (C) Fluorescent micrographs of the replated partially differentiated hMSC stained with FITCPhalloidin (green) and DAPI (blue). The cells were initially cultured on implant and control substrates. They were then trypsinized and replated on tissue culture plate. Imaging was done after 3days of replating. (D) Alizarin red staining of the replated partially differentiated hMSC. Imaging was done after 3 days of replating. ‘*’ designates statistical significance with p < 0.05.

3.7. In vivo assessment of the performance of the growth factor loaded implant In vivo analysis showed that the implants were capable of restoring their structural integrity. From Figure 10 A and 10 B, it is evident that the implants were intact and mechanically maneuverable. One noticeable point is that none of the implants (with or without VEGF) got dried after implantation (semi-translucent appearance confirmed the wet condition). This clearly 30 ACS Paragon Plus Environment

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suggests that there was an effective exchange of fluids in vivo. A critical analysis further confirmed the existence of a cohesive interaction between the native tissue and implant (marked as (2)) which can be considered as a kind of tissue integration. Moreover, during the post operative period none of the mice died nor showed any kind of discomfort. These facts altogether suggests about its potential as an implant. Another prime goal of this experiment was to check whether VEGF loaded implant could induce angiogenesis in vivo or not. Visual inspection of the area that was in contact with the implants clearly showed the existence prominent vasculature in case of VEGF loaded implant (Figure 10 B2) which was absent in case of implant without growth factor (Figure 10 B1). The rational explanation of such outcome is the prolonged release of VEGF from the implant which activates the endothelial cells at the release site resulting in neovascularization. This result strongly favors the applicability of the implant.



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Figure 10 In vivo study of the angiogenic property of VEGF loaded implant. Control implant (without VEGF) and implant loaded with VEGF were implanted at the left and right subcutaneous dorsal pockets of healthy Swiss albino mice. Mice were sacrificed after 7 days of implantation and the subcutaneous area that was in contact with implant was investigated. (A1) implant without VEGF and (A2) implant with VEGF. [1] Implant, [2] Cohesive interaction between the native tissue and implant, [3] tissue area that was in contact with the implants, [4] blood vessel. Enlarge view of tissue area that was in contact with the implants. (B1) implant without VEGF, (B2) implant with VEGF.

4. Conclusion The present study delineates the fabrication of a novel kind of micro-patterned bone implant, capable of delivering therapeutics in a spatio-temporally controlled manner. We prepared the implant by stacking 2D HCP layer of alginate beads. Due to the intrinsic structural feature of HCP, there was formation of an array of tetrahedral and octahedral pores of definite size and geometry inside the implants, which made the interior micro-patterned. One crucial part of this scaffolding technique is that, though it generates micropores, but no microscale-tool is required to achieve the desired interior. Moreover, the size of the pores could be controlled by simply modifying the dimension of the alginate beads. Mechanical analysis of the implant showed an increase in the compressive modulus upon incubation in SBF. Such property will be beneficial for its in vivo application. However, the compressive modulus was found many fold less than that of non-load bearing bone. This is a common problem associated with most of the hydrogels for bone tissue engineering applications. This problem could be surmounted by incorporation of reinforcing materials like nano-ceramics or carbon based nano-materials (carbon nanotube, graphene oxide). By conducting the diffusion study on a porous solid agar plate, we showed that spatio-temporal tuning of release kinetics is possible by changing the coordinates of depot (drug loaded beads) inside the implant. As different beads can be loaded with different drugs, so this bead based implant can further be explored for simultaneous delivery of multiple drugs in a spatio-temporally controlled fashion. The analysis of the biological performances of the implant confirmed its osteoconductive nature. A high expression of VEGF by the bone cells cultured on the scaffold further implicated its beneficial role in angiogenesis, a requisite for successful bone tissue engineering. Osteogenic property of the implant was further confirmed by the higher 32 ACS Paragon Plus Environment

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expression of cellular differentiation markers like ALP, Runx2 and Col1 in hMSC. Finally, we showed that showed that the VEGF loaded implant was capable of inducing angiogenesis in vivo in mice. In conclusion, we have established the bead based implant as novel prototype for bone tissue engineering.

Acknowledgement Authors would like to extend sincere thank to Prof. S. K. Bhutia, NIT Rourkela for extending fluorescence imaging facility.

Supporting Information Regression coefficient (R2 ) of different drug release kinetics model.

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Graphical Abstract


Alginate Bead Based Hexagonal Close Packed 3D Implant for Bone

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