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Article Cite This: ACS Omega 2019, 4, 1504−1515
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Regulation of Human Osteoblast-to-Osteocyte Differentiation by Direct-Write 3D Microperiodic Hydroxyapatite Scaffolds Kulwinder Kaur, Sanskrita Das, and Sourabh Ghosh*
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Regenerative Engineering Laboratory, Department of Textile Technology, Indian Institute of Technology Delhi, New Delhi 110016, India ABSTRACT: Culturing osteocytes on planer surfaces on Petri dish in vitro fails to recapitulate their natural orientation as well as biological functionality in human bone. Ability to recapitulate spatial arrangement of osteoblasts can govern signaling cascades for osteoblast-to-osteocyte differentiation and osteogenesis. Therefore, in this study, we explored how direct-write microperiodic hydroxyapatite (HAP) threedimensional (3D) scaffolds can modulate osteocyte differentiation as compared to two-dimensional (2D) HAP scaffolds. Increased level of osteocalcin expression and relatively early mineralization on 2D HAP were indicative of osteoblast maturation. On the other hand, unique features of 3D HAP direct-write scaffolds, in terms of the architecture and subtle difference in chemical composition, surface roughness, and stiffness, play a pivotal role in governing the complex mechanism of differentiation of osteoblast to osteocyte-like phenotype. This was characterized by sequential events involving modulation in proliferation rate, time-dependent downregulation in the expression of osteoblast markers, collagen type I and alkaline phosphatase, followed by the unregulated expression of matrix metalloproteinase, osteocytic terminal differentiation markers (podoplanin, sclerostin), and dendritic extension formation. Thus, establishment of a simple in vitro model to study osteoblast differentiation with respect to specific cues would offer potential as an investigative platform and versatile tool for developing a 3D in vitro model for regenerative medicine.
1. INTRODUCTION Human bones are perpetually remodeled throughout life. Bone mineralization takes place in two stages. At first stage, it is coordinated by osteoblasts and subsequent differentiation of osteoblasts to osteocytes. During the mineralization process, osteoblastic cells undergo interactions with various inorganic ions,1 proteins,2 and calcium phosphate-enriched surface.3,4 Secondary phase of mineralization is regulated through crystal maturation and matrix remodeling by the osteocytic network.5 Osteocytes represent the most abundant (90−95%) cell population of adult bone, embedded within the mineralized extracellular matrix (“lacunae”), express cytoplasmic dendritic extensions which penetrate throughout the bone canaliculi, whose primary function is to regulate bone homeostasis by sensing mechanical signals,6 and convert these cues to biochemical signals and communication with the osteoclasts and osteoblasts by means of signaling mechanism to remodel bone.7 Osteoprogenitors differentiate to their mature phenotype in the sequence of differentiation events via preosteoblasts and osteoblasts and ultimately form osteocytes. It is still not clear whether terminally differentiated osteocytes can dedifferentiate to osteoblasts8 or to what extent osteoblast-toosteocyte differentiation can be modulated by the biomaterials.9 To be able to explain the biomaterial-induced, mechanotransduction-mediated osteocytic lineage differentiation, detailed understanding of the optimal physicochemical characteristics of the underlying scaffolds, such as dimension© 2019 American Chemical Society
ality, geometry, surface chemistry, roughness, and stiffness, would be of great interest. Establishing a relevant in vitro model to investigate osteoblast-to-osteocyte differentiation would offer potential as an investigative platform to study osteoblast differentiation with respect to specific cues. There is paucity of in vitro studies on osteocytic differentiation because of several challenges. First, there is a difficulty in isolation of primary osteocytes that are deeply embedded within the hard bone tissue. Second, terminally differentiated primary osteocytes cannot be expanded in vitro in monolayer without loss of phenotype.10 Third, few osteocyte cell lines have been established from mouse origin or osteosarcoma tissues, which do not fully capture the phenotypic features of human mature osteocytes.11,12 Genetically manipulating such primary cells into immortalized cell lines modifies the cells’ gene expression profiles and has serious implications on the physical integrity of the structure and function of genome. Fourth, the expression of important markers of mature osteocytes, such as fibroblast growth factor 23 and sclerostin (SOST), is heavily compromised in some of these cell lines, during conventional monolayer culture.1,13 In an attempt to simulate the in vivo microenvironment for osteoblast-to-osteocyte differentiation, osteocyte growth, and Received: November 23, 2018 Accepted: January 2, 2019 Published: January 16, 2019 1504
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Figure 1. Rheological characterization of HAP ink. (a) Viscosity at varied shear rates and (b) elastic and viscous modulus.
events from osteoblast to osteocyte-like phenotype have been well reported in monolayer culture,15 the underlying intrinsic mechanisms involved in the translational stages and the role of cell−matrix interactions in the process remain elusive.16 Structural organization of osteoblasts as a function of modulation in underlying matrix mechanics, such as stiffness and surface roughness, is found to affect cellular behaviors such as attachment, proliferation, migration, and differentiation.17 In the present work, we have attempted to study the maturation and differentiation of primary human osteoblasts using a direct-write 3D printed hydroxyapatite (HAP)-based in vitro model system. A comparative evaluation was made on osteoblast behavior between 2D and 3D printed HAP scaffolds. Unlike the conventional methods of random porous scaffold manufacturing, direct-write 3D printing is capable to fabricate advanced scaffolds with precisely controlled architecture and porosity,18,19 thereby simulating the exact 3D microenvironment of the native tissue and guiding the orientation of the seeded cells. We hypothesized that the osteoconductive property of HAP would support cellular attachment and differentiation on both 2D and 3D structures; however, different geometries may modulate cellular morphology that will further influence the terminal differentiation of osteoblasts. Interestingly, we observed that even subtle difference in materials chemistry, stiffness, and surface roughness can have a significant impact on the proliferation, differentiation kinetics, and matrix remodeling of seeded primary osteoblast cells.
developmental processes, collagen type I hydrogel-based in vitro models were developed, where the primary preosteoblast cells were embedded within a dense fibrillar collagenous substrate to simulate the density and stiffness of the osteoid bone.2 However, enhanced osteocytic differentiation of MC3T3-E1 cells was noticed on the soft collagen-based matrix.3 MC3T3-E1 murine osteoblasts grown over collagen type I gel differentiated into osteocytic cells with increased osteoblast and osteocyte-specific mRNA expression unlike cells cultured in monolayer4,5 or collagen type I-coated surface.6,7 In another study, human osteosarcoma SaOS2 cell line was cultured on collagen type I gel for up to 35 days under mineralizing conditions. Results revealed the synthesis of a mineralized matrix with the presence of mature osteocytes, which was characterized by the gene expression profile [upregulation of dentin matrix protein 1 (DMP1), SOST, and downregulation of RUNX2 and collagen type I genes] and typical dendritic morphology.8 However, collagen gel as a three-dimensional (3D) matrix possesses poor mechanical properties and suffers from loss of shape and consistency during the culture period because of its inherent tendency to contract, resulting in reduced porosity of the scaffold, which in turn can distort cell morphology and affect the resultant cellular behavior.9 Moreover, the above-mentioned studies have used osteoblastic cell lines, and the expression of podoplanin, an important marker for early osteocyte differentiation, could not be seen. Another important aspect is to replicate the complexity of 3D architectures to support the 3D networked osteocytes in vitro, which are typically aligned along concentric lamellae in human bone.14 Human primary osteoblasts cultured over 40− 80 μm biphasic calcium phosphate particles12 differentiated toward osteocyte-like phenotype with subsequent downregulation of osteoblast-specific markers, in comparison to their two-dimensional (2D) counterparts. However, these 3D cultures based on aggregates of calcium phosphate particles failed to reproduce the typical 3D ordered osteocyte network because of the formation of cellular aggregates randomly dispersed among interstitial spaces of calcium phosphate particles. Because of lack of appropriate in vitro 3D models, it still remains unclear that how osteoblast cells get entrapped within the mineralized matrix and how these osteocytes further differentiate and mature into cell−matrix and cell−cell interactions lead to the development of the mechanically remodeled, functional bone tissue. Although the cascades of
2. RESULTS AND DISCUSSION 2.1. Structure Analysis. The viscosity of HAP suspension at 25 °C was noticed to be 40.3 Pa s at the shear rate of 10 s−1, which was decreased to 9.1 and 2.2 Pa s at shear rates of 100 and 1000 s−1, respectively (Figure 1a), thereby displaying a shear thinning behavior. At 1% strain and 1 Hz frequency, the elastic modulus and viscous modulus of HAP suspension were observed to be 7.9 × 105 and 1.27 × 105 Pa, respectively (Figure 1b). Hence, below 100% strain, the shear thinning nature of HAP ink and higher elastic modulus than viscous modulus demonstrated the fidelity of fabricating selfsupportive scaffolds without significant deformation.20 HAP scaffolds fabricated in this study are an advancement in comparison to earlier studies21 because of the use of a cellulose derivative as a thickening agent. Furthermore, longer time 1505
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vibrations of PO43− units. Bands near ∼1400 and ∼2800 cm−1 were due to the stretching vibrations of CO32− groups. The band at wavelength 871 cm−1 was due to HPO42− groups. These bands were more predominant in the 3D scaffold. The nonapatitic species (HPO42− and CO32−) could be easily modified according to the desired composition and also these groups are present at higher concentrations in newly formed bone crystals.23 Apatite crystals containing nonapatitic ionic groups play a crucial role in differentiation of osteoblast cells.22,24 In the case of 3D HAP scaffolds, vibrations corresponding to HPO42− and PO43− were more intense as compared to the 2D and powder sample so that the 3D scaffold would be more reactive toward osteoblast differentiation as compared to the 2D or powder sample. These findings validate the XRD results that the geometry and materials chemistry of the 3D HAP scaffold would promote osteoblast differentiation. In Figure 3, vibrations at ∼556 and
duration for sonication resulted in uniform dispersion and disintegration of HAP particles. X-ray diffraction (XRD) diffractograms of powdered HAP and sintered 2D and 3D scaffolds were found to be crystalline in nature (Figure 2A). Distinct peaks at 31.4, 32.4, 32.5, 38.3,
Figure 2. (A) XRD diffractograms (■ HAP, ⊙ calcium carbonate) and (B) FTIR spectra of calcinated HAP powder, 2D HAP, and 3D HAP.
and 44.5° confirmed the presence of calcium carbonate phase with Joint Committee on Powder Diffraction Standards (JCPDS) card number 071-2392 and peaks at 25.8, 31.8, and 32.9° confirmed the presence of HAP phase with JCPDS card number 086-1203 confirmed by Pcpdfwin software. The peak corresponding to the HAP phase at 31.8° displayed higher intensity in 3D HAP scaffold diffractogram as compared to 2D and powder sample diffractograms (inset of Figure 2A). Higher content of the HAP phase, that is, phosphate contents, was preserved better in 3D geometry as compared to that in 2D geometry after sintering because of homogenized sintering in 3D HAP scaffolds compared to that in 2D geometry. Because of the presence of higher content of phosphate and lower content of carbonate in a 3D scaffold, it would be more inductive for osteoblast differentiation as compared to 2D, akin to in vivo condition promoting bone formation.22 Attenuated total reflection−Fourier transform infrared (ATR−FTIR) spectra for all the samples are presented in Figure 2B. Intense bands in all the samples were in the range of 1010−1090 cm−1, which represented the antisymmetric
Figure 3. Representative SEM micrographs: (a) calcinated powder HAP, (b) sintered 2D HAP, (c) CAD image, optical image of 3D HAP scaffolds (d) top view and (e) side view, (f) SEM micrograph of 3D HAP, and (g,h) magnified SEM images of 3D HAP.
∼602 cm−1 are fingerprint vibrations of HAP.25,26 Vibrations at ∼630 and ∼3600 cm−1 in all the samples correspond to [OH]− because of the presence of OH group in the HAP matrix.27,28 Difference in surface properties of 2D and 3D HAP scaffolds after sintering was clearly distinguishable (Figure 3). The scanning electron microscopy (SEM) micrographs revealed that the sintering temperature caused characteristic grains in 1506
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roughness (Rms) values for 2D and 3D scaffolds were 8.43 ± 0.54 and 61 ± 1.71, respectively, and stiffness values were 296 ± 12.3 and 459 ± 22.4 Pa as calculated by using eqs 1 and 2, respectively. In the first step after implantation, development of an interface between the bone and the implant material depends upon the interaction between the osteoblast and the material.30 Adhesion of osteoblast cells on the surface of a material is an extremely important phenomenon for the first bone−material interaction. Highly rough and less stiff surfaces promote the osteogenesis at early stages of bone healing.31−33 Different methods have been developed to make rough implant surfaces for the betterment of clinical performance of the implant material,34,35 such as etching of titanium implants with strong acids has been reported.30 Here, we report the effect of geometry on the roughness and stiffness of the sample and we found that 3D HAP scaffolds are highly rough and less stiff as compared to the 2D HAP sample, which lead to the high rate of osteoblast cell adhesion as compared to the 2D HAP sample. Results of AFM analysis validated the results of XRD, ATR−FTIR, and SEM that 3D HAP scaffolds were more reactive and favorable substrates for osteoblast differentiation to osteocyte. Taken together, from all the physical characterizations of 2D and 3D HAP scaffolds, it was concluded that the geometry, dimensionality, materials chemistry, surface roughness, and stiffness of the 3D scaffold would be favorable substrates for osteoblast differentiation to osteocyte as compared to the 2D HAP sample. 2.2. SEM Morphological Responses of Osteoblasts on HAP. Osteoblast cells were well attached and displayed
3D scaffolds, akin to earlier reports29 (Figure 3h). Grains were not prominent in the 2D HAP sample (Figure 3b). This was due to the presence of higher content of calcium carbonate in the 2D sample, as the melting point of calcium carbonate is 850 °C, which is lower than the sintering temperature of samples, that is, 1250 °C. Melting of calcium carbonate phase leads to disappearance of grains in 2D samples. While in the case of 3D HAP scaffolds, phosphate is present at higher content and the melting temperature of HAP is 1700 °C, which is higher than the sintering temperature, so clear grains were observed in 3D HAP scaffolds. Therefore, results of SEM analysis also validate the results of XRD and ATR−FTIR analysis. Surface properties of an implantable biomaterial have determinant effects on the biological behavior. The surface morphology of prepared scaffolds was observed by atomic force microscopy (AFM) to measure the roughness and stiffness of 2D and 3D HAP scaffolds (Figure 4). Surface
Figure 4. AFM images of (a) 2D HAP and (b) 3D HAP to elucidate difference in surface morphology.
Figure 5. Cellular morphology of osteoblast on 2D HAP and 3D HAP scaffolds. Week 1 constructs: (A,B), week 4 constructs: (C−F), week 6 constructs: (G,H). I is the magnified image of differentiated osteoblast on 3D HAP week 6, respectively. 1507
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Figure 6. Cellular responses within 2D HAP and 3D HAP: (A) DNA contents, (B) ALP, and (C) total collagen content (ng/ng of DNA) from the printed constructs in osteogenic condition (# and * are significant between 2D HAP and 3D HAP).
due to the embedding of the osteoblasts within the calcium phosphate particles, which presented an osteoid-like environment to the cells. However, a major concern for such type of 3D cultures is that cellular entrapment within particles may lead to cell aggregation, thereby hindering the transition of osteoblast to osteocyte-like phenotype. Moreover, in the above-mentioned study, stellate-shaped osteocyte-like cells with characteristic dendritic features were not evident. On the other hand, osteoblasts cultured on 2D HAP scaffolds showed early evidence of mineralization as compared to 3D HAP scaffolds (Figure 5C,E,G). This differential pattern of mineralization on the two HAP surfaces highlights the importance of scaffold geometry on osteoblast differentiation. As concluded from all the physical characterizations explained in Section 2.1, 3D scaffolds were more reactive to induce osteoblast differentiation because of the presence of higher phosphate content and lower stiffness value. These factors contributed to more pronounced osteocyte differentiation in 3D HAP scaffolds as compared to the 2D HAP sample. 2.3. Evaluation of DNA Content and Biochemical Assay. No significant difference could be observed in DNA content between 2D HAP and 3D HAP scaffolds on 2nd week {Figure 6A}. However, the DNA content was observed to be significantly different between the two conditions on 4th (2D HAP and 3D HAP, p = 0.008) and 6th week (2D HAP and 3D HAP, p = 0.02), respectively. Also, significant increase in DNA content could be observed from 2nd to 4th week (p = 0.002 and p = 0.00006 for 2D HAP and 3D HAP, respectively). Further, the increased DNA levels (1.07- and 1.06-fold on 4th and 6th week) for 3D HAP over 2D HAP {Figure 6A} highlighted the importance of 3D geometry with interconnected pores, which is required for promoting attachment, proliferation, and migration of osteoblasts, as well as facilitating the transport of nutrients and metabolic wastes. After 6 weeks of culture in differentiation media, the amount of alkaline phosphatase (ALP) released by osteoblasts was highest for 3D HAP scaffolds (15.2 IU/L) than that of 2D HAP (9.3 IU/L) {Figure 6B}. The ALP level was estimated to
osteoblastic morphology on all the substrates, that is, 2D HAP (Figure 5A) and 3D HAP (Figure 5B) within first 1 week. However, modulation in their morphology and mineralizing tendency could be observed with increasing culture time points. On 4th week, crystal-like substances or mineralized deposits could be observed on both HAP scaffolds, albeit more in 2D HAP scaffolds (Figure 5C,E). On 3D HAP scaffolds, osteoblasts displayed extensive cytoplasmic dendritic structures (Figure 5D,F), characteristic features of osteocytes, which were absent in cells grown on 2D HAP scaffolds. The osteocytes visualized in the field of view formed extended dendrites that connected not only the neighboring osteocytes but formed an extensive interconnected network that extended toward the interfilament spaces across the 3D printed scaffold. This characteristic interwoven organization of the dense dendritic canalicular network exhibited close resemblance to the in vivo bone physiology.36 On 6th week, extensive mineralization could be observed on 2D HAP (Figure 5G). Extensive dendritic features with stellate morphology, another hallmark of osteocytes, along with mineral deposits, could be observed on 3D HAP, which was completely missing on 2D HAP (Figure 5H,I). Taken together, although the same material was used to fabricate both the structures, osteoblasts displayed distinct differences in their response to the scaffolds of dissimilar geometry. Osteoblasts cultured on 3D HAP had produced a cellular network with an extensive filamentous structure (Figure 5D,F,H,I), closely resembling the canaliculi-like ultrastructural characteristics of osteocytes. This observation concurs with the earlier study12 where human primary osteoblasts seeded within the interspace of calcium phosphate granules have exhibited the characteristics of osteocyte-like cells. However, distinct differences do exist between the two studies in terms of composition and source of the material, culture duration, precise 3D geometry of the scaffold, and culture method. Unlike our study, Boukhechba et al.12 reported osteocytic differentiation of osteoblasts just after 1 week of culture on calcium phosphate particles. This could be 1508
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Figure 7. Evaluation of gene expression by RT-PCR: (A) ALP, (B) RUNX2, (C) collagen type 1, (D) osterix, (E) osteonectin, (F) osteopontin, (G) β-catenin, (H) BSP-2, (I) osteocalcin, (J) MMP-2, (K) podoplanin, and (L) SOST. Data expressed as relative fold change for specific gene expression in 2D HAP and 3D HAP constructs at the time point mentioned (# and * are significant between 2D HAP and 3D HAP).
expression of osteocalcin and collagen type I.38,39 The expression of RUNX2, initial marker of the osteogenic cell lineage, demonstrated drastic upregulation in 4 weeks. Expression of ALP on 2nd week (p < 0.01) and RUNX2 on 2nd and 4th week (p < 0.01) was observed to be significantly higher in the case of 2D HAP scaffolds as compared to that in 3D HAP scaffolds {Figure 7A,B}. The level of ALP on the 2nd week was 2.4-fold upregulated and that of RUNX2 on 2nd and 4th week was 1.4- and 1.3-fold upregulated in 2D HAP scaffolds as compared to that in 3D HAP scaffolds. However, the ALP level was 2.7- and 21.5-fold upregulated on 4th and 6th week, and the RUNX2 level was 75.3-fold upregulated in 3D HAP scaffolds than that in 2D HAP scaffolds. Similarly, the levels of ALP and RUNX2 were observed to be 1.5-, 2.7-, and 2.4-fold and 1.4-, 1.6-, and 1.2-fold upregulated in 3D HAP scaffolds because of osteogenic differentiation owing to the presence of higher content of phosphate ions in 3D HAP scaffolds as compared to that in 2D HAP because of higher surface roughness and lower stiffness of 3D HAP scaffolds, as lower stiffness favors osteoblast differentiation.16 Comparing between 2D and 3D HAP scaffolds, the level of collagen type I gene expression was upregulated with 1.2-fold in 2D HAP on the 2nd week, whereas on 4th and 6th week, the level was downregulated by 5 and 29 folds, whereas in the case
be significantly higher in the case of 3D HAP than that of 2D HAP scaffolds on 4th (15.2 IU/L and 9.3 IU/L, p = 0.001) and 6th (12.9 IU/L and 6.9 IU/L, p = 0.008) week, respectively. Higher value of ALP content might be due to the presence of higher content of the phosphate group in 3D HAP scaffolds as confirmed from XRD and ATR−FTIR. Significant increase in collagen production could be observed in 3D HAP scaffolds on 4th week (p = 0.03), followed by decrease in 6th week (p = 0.002) {Figure 6C}. 2.4. Gene Expression Analysis. The differentiation and commitment of osteoblasts are controlled by complex activities of gene expression such as transcriptional and transduction regulation of genes. Therefore, in order to elucidate the effect of scaffold dimensionality and to establish an in vitro differentiation model system, we implicated different signal pathways and transcription factors for osteoblast differentiation on 2D HAP and 3D HAP in a network manner (Figure 7). 2.4.1. Osteoblast Markers. ALP, RUNX2, collagen type I, osterix, and BSP are early markers of osteoblast differentiation.37 Osteonectin is a midstage marker, whereas osteocalcin and osteopontin are late stage markers. ALP and RUNX2 (core binding factor alpha 1) are early markers and key transcription factors for osteoblastic differentiation. For osteoblast differentiation, RUNX2 is crucial as it activates the 1509
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reasonable starting point to study the intricate transition of osteoblast-to-osteocyte differentiation in 3D HAP scaffolds. 2.5. Alizarin Red Staining and Immunofluorescence Analysis. In order to validate gene expression data, alizarin red staining and immunofluorescence analysis were performed on 2D and 3D constructs for assessing the presence of mineralized nodules and production of collagen type I. Intense alizarin red staining on 2D HAP scaffolds indicates that cells grown on 2D geometry underwent more extensive mineralization during the later stages of osteogenesis than cells on 3D HAP scaffolds (Figure 8A,B). Immunofluorescence staining
of 3D HAP scaffolds, collagen type I gene was upregulated by 14 folds on the 4th week and then downregulated by 16 folds on the 6th week because of temporal osteogenic differentiation40 {Figure 7C}. No significant difference could be observed between both these scaffolds on 4th week. Osterix, a downstream target of RUNX2,41 was upregulated by 150 folds from 2nd to 4th week and downregulated to 8 folds on 6th week in 2D HAP. In the case of 3D HAP scaffolds, osterix was upregulated by 160 folds on 4th week and by 110 folds on 6th week {Figure 7D}. Osteonectin is a glycoprotein synthesized abundantly by the cells of the osteoblastic lineage,10 and osteopontin is a cell- and HAP-binding protein.42 These midmarker proteins along with collagen type I are critically important for matrix mineralization.43 Expression pattern of osteonectin and osteopontin has elevated in 3D HAP scaffolds by 1.8-, 1.8-, and 50.8-fold and 8.8-, 1.9-, and 281-fold as compared to 2D HAP at all the time points {Figure 7E,F}. Similarly, the expression pattern of β-catenin, a cytoplasmic phosphoprotein under resting conditions and the key mediator of the Wnt signaling pathway,44 was observed to be 2-fold upregulated in 3D HAP scaffolds than that in 2D HAP scaffolds by 4th week {Figure 7G}. Bone sialoprotein-2 (BSP-2), which is mainly involved in the formation of apatite crystal,41 was found to be 2- and 8.5-fold higher in 3D HAP scaffolds {Figure 7H}. In other conditions, significant difference could not be observed. Osteocalcin is a noncollagenous calcium-binding protein, which tightly binds onto the HA surface and is indicative of late stage osteoblast differentiation.41 On 4th week, cells from 2D HAP scaffolds have displayed significant upregulation with 1.3-fold increase as compared to 3D HAP scaffolds {Figure 7I}. However, by 6th week, expression of osteocalcin has drastically reduced in the case of 2D HAP scaffolds with 15.8-fold upregulation in 3D HAP scaffolds. Additionally, matrix metalloproteinase-2 (MMP-2) activity is required for the osteoblast-to-osteocyte differentiation, based on the activation of latent TGF-β entrapped within the surrounding matrix.45 Specifically, cells harvested from 2D HAP scaffolds have expressed 1.5-fold upregulation of MMP-2 as compared to 3D HAP scaffolds by 4th week {Figure 7J}. 2.4.2. Osteocyte Markers. Interestingly, podoplanin, which is an early marker for young osteocytes,46 has displayed significantly elevated expression (p < 0.01) by 4th week in 3D HAP scaffolds with 1.5-fold upregulation as compared to that in 2D HAP scaffolds {Figure 7K}, but its expression was drastically decreased on 6th week. This probably suggests that podoplanin is probably crucial for the initial development of dendrites but not for the subsequent maintenance of dendritic morphology. Indeed, podoplanin expression was noticed predominantly in the embedded cells.47 Some other studies also showed downregulation of podoplanin expression at later stages of osteocytic differentiation.42 SOST was observed to be 4.8-fold upregulated than that of 2D HAP scaffolds on 4th week {Figure 7L}. Importantly, we could not observe any SOST expression on 2D HAP scaffolds on 6th week. The SOST protein, a secreted WNT/β-catenin antagonist, inhibits Wnt signaling and subsequently inhibits bone formation, which is probably the reason for decrease in the osteogenic expression markers by 6th week, putatively showing features of bone homoeostasis.48,49 Interestingly, no DMP-1 expression could be observed at any time points. Altogether, gene expression study provided some confidence that this in vitro culture condition might be a
Figure 8. Alizarin red staining showing mineralized deposits: (A) 2D HAP and (B) 3D HAP. Immunofluorescence staining for type 1 collagen: (C) 2D HAP and (D) 3D HAP; phalloidin staining for actin filaments: (E) 2D HAP and (F) 3D HAP. (G) Yellow arrow showing the presence of dendritic cytoplasmic processes only in 3D HAP constructs.
showed punctuated morphology of collagen throughout the cellular cytoplasm in the 2D matrix, whereas cells on 3D HAP scaffolds demonstrated more organized collagen fibrils, albeit in intracellular regions (Figure 8C,D). There was a drastic difference in the pattern of actin cytoskeleton expression in cells between 2D and 3D constructs. Extended cytoplasmic actin fibers were observed evidently on cell cytoplasm as well as, more distinctly, actin bundles were visible in cellular extensions in 3D HAP constructs, compared to the cells in 2D HAP constructs (Figure 8E−G), depicting definitive evidence of transition from osteoblast to osteocytes. 2.6. Insights Generated from the Osteoblast-toOsteocyte Differentiation Model. Various studies have explored HAP scaffolds in several forms (2D, randomly porous 1510
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Second, presence of higher content of phosphate ions in 3D HAP scaffolds as compared to 2D HAP after sintering may be responsible for enhancing the osteoblast differentiation and cell transition to osteocyte. Phosphate ions enter into the cell through the normal transporter mechanism and promote the ALP activity and stimulate the osteoblast differentiation. Sai et al.24 studied the capacity of calcium phosphate such as HAP, βtricalcium phosphate, and octacalcium phosphate (OCP) to promote osteoblast-to-osteocyte differentiation in vitro after 35 days. They observed that on immersion of OCP into the media, hydrolysis of the OCP occurs, which released phosphate ions into the media by the reaction57
or 3D printed), without proper scientific justifications how such morphologies may modulate osteogenic or osteocytogenic signaling pathways. This led to diverse outcome and poor reproducibility of osteogenesis.50 Early occurrence of osteocytogenesis would leave insufficient time for the production of bone matrix; at the same time, delayed mineralization would cause poor mechanical strength of the engineered constructs and in turn would result in disappointing clinical outcome. This study highlights importance to evaluate the ideal scaffold architecture for optimized osteocytogenesis, which may offer potential to mitigate the heterogeneity of bone regeneration because of patient-to-patient variation. In the present study, human primary osteoblasts-seeded 2D HAP constructs revealed evidence of early mineralization with extensive mineral deposition, along with the increased level of osteocalcin that is indicative of osteoblast maturation. On the other hand, 3D HAP constructs showed the formation of canaliculi-like structures, overall enhanced osteogenic gene expression, as well as temporal expression of podoplanin and final upregulation of SOST gene expression, indicative of osteocytic differentiation. So far, the osteoblast−osteocyte differentiation has been studied in the context of exogenously added factors such as oncostatin M51 or retinoic acid.52 For example, Mattinzoli et al.53 reported that the addition of 5 mM retinoic acid could differentiate both MC3T3-E1 osteoblasts and primary calvarial cells to the osteocyte lineage in 10 days. However, unique finding of the present study is the scaffold architecture that may modulate the osteoblast−osteocyte differentiation. A constellation of factors might be responsible for the differential behavior of osteoblasts over 2D and 3D HAP scaffolds prepared by direct-write assembly (DWA). First, culturing cells on 2D surfaces induces the osteoblast polarization, leading to a flattened and spheroidal cell shape, as well as it may not allow for sufficient multilayering of cells to translate the controlled organization in multiple layers of cells, thus accurately reproducing their natural morphological conformation.47 As opposed to this, 3D printed scaffolds provided a guided structure for the cells to allow optimal spreading area, with aligned organizations that more accurately reproduce the native architecture of bone and hence may involve mediators of cell−cell contact, such as gap junction proteins. This highlights one of the prime reasons for the development of microperiodic scaffolds for bone tissue engineering applications. In our study, we reported the increased levels of β-catenin, a key component of the canonical Wnt pathway, which also stimulates connexin 43 expression, a major component of gap junctions and hemichannels between osteocytes.54 Hence, the increased expression of β-catenin and decreased MMP-2 activity suggests that at least a part of the enhanced osteogenic signal in 3D HAP scaffolds may be due to gap junction-mediated cell communication. Studying the expression of other classical gap junction proteins (such as connexin 43)55 may provide useful insights about the mechanism behind the terminal differentiation of osteoblast to osteocyte-like phenotype in 3D HAP scaffolds. TanakaKamioka et al.56 showed several pieces of evidence of focal convergences of actin fibril bundles with the dendritic extensions in osteocytes, which provide the mechanosensory capacity of osteocytes. Our findings clearly showed evidence of bundled actin cytoskeletal organization in 3D cell constructs, providing some confidence about the success of establishment of the in vitro osteoblast-to-osteocyte differentiation model.
10 Ca8H 2(PO4 )6 ·5H 2O 8 → Ca10(PO4 )6 (OH)2 +
3 17 H3PO4 + H 2O 2 4
They found that OCP has the highest capacity to differentiate the IDG-SW3 cell line because of the presence of higher concentration of phosphate ions in OCP released during hydrolysis. These phosphate ions will transport into cells and enhance the ALP peak in the cells. SOST/sclerostin and FGF23 gene expression were also upregulated after 35 days for OCP samples. Third, substrate stiffness and roughness are an important mediator of osteoblastic differentiation. Higher content of HAP in 3D HAP is due to homogenized sintering of 3D HAP, compared to 2D HAP, which in turn caused lower stiffness and higher roughness of the 3D HAP scaffold. Enhanced Wnt/βcatenin signaling pathways and osteocytic marker expressions indicated that osteoblasts could sense this subtle difference in matrix stiffness and surface roughness to translate that cue into biochemical signals to affect differentiation to osteocytes and eventually modulate mineralization and matrix remodeling. It would be interesting to explore how closely one may recapitulate the architectural complexity of lacunar−canalicular topography present in the osteon in human bone by 3D printing and how closely HAP scaffolds may be made by recapitulating the architectures of woven and lamellar bone so that the osteocytes are completely embedded as well as cellular shape and orientation are controlled to govern osteoblast-toosteocyte differentiation. This insight will help to develop rationally designed, better engineered bone graft, as well as to check the efficacy of new drug molecules for various osteopathologic disorders. In future, further improved in vitro 3D printed model for human osteoblast-to-osteocyte differentiation will be developed by the application of fluidflow shear stress on this established model in an attempt to improve the cell communication networking for developing dendritic structures. In addition to the scaffold architecture and chemistry, rational mixing of several soluble factors in appropriate combinations such as vitamin D3, estrogen, TGFβ, thyroid hormone, and/or cytokines26 in varied ratio may elucidate indispensable role and synergistic effects of those components in the differentiation pathway of osteoblast to osteocyte.
3. CONCLUSIONS Our study provides an experimental insight toward the establishment of a 3D printed in vitro model system for studying osteoblast-to-osteocytic differentiation with respect to varied geometry, dimensionality, surface roughness, stiffness, 1511
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spacing of 600 μm with a total of six consecutive layers. Subsequent layers were deposited at a 90° angle to the underlying layer to fabricate a 3D scaffold. 2D HAP scaffolds were fabricated by depositing the HAP ink onto the flat glass surface, which was dried at 60 °C. Afterward, the dried HAP scaffolds were moved from the glass surface and were turned upside down to obtain the flat 2D geometry. The overall dimension for 2D HAP in the x−y plane was kept similar to that of 3D scaffolds. Further, the 2D and 3D HAP scaffolds were sintered at 1250 °C for densification without decomposition.61 Specifically, the scaffolds were heated at 1 °C/min to 400 °C for 1 h, then at 3 °C/min to 900 °C for 2 h, and, finally, at 4 °C/min to the sintering temperature with a holding time of 2 h. After printing, the resultant diameter for 3D HAP was 266 ± 17 μm, whereas the structural deviation from the set dimensions was approximately 2.4%. 4.4. Physical and Biological Characterizations. 4.4.1. Rheological Measurements. Prepared printable ink was physically characterized by a rheometer MCR 302, Anton Paar with a cone-and-plate geometry (cone diameter of 25 mm with 1° angle) at 25 °C using the environmental cuff. To predict printability, the viscosity of the suspension was measured at different shear rates in the range from 0.1 to 1000 s−1. The dynamic elastic modulus (G′) and viscous modulus (G″) were also measured at 25 °C in an oscillatory mode. In the amplitude sweep measurement, the HAP ink was tested for the strain value ranging from 0.1 to 104% with a frequency constant at 1 Hz in the frequency sweep measurement, and the HAP ink was tested for the frequency ranging from 0.1 to 100 Hz with a strain constant at 5%. Samples were taken in triplicates. 4.4.2. AFM Nano Indentations. AFM (Dimension ICON with Scan Asyst, Bruker) study was done in tapping mode using a rotated monolithic silicon tip (Tap300Al-G, Budget Probes, Bulgaria) to measure the root-mean-square roughness (Rms) of the 2D and 3D HAP scaffolds. The elastic modulus of samples (n = 5/group) was measured using AFM operated in force mode. First, the sample was inspected by AFM using the tapping mode to locate the scaffold area for indentation. Next, the AFM was switched to force mode, and indentation was done. In different regions, force−distance curves for each sample were measured, the calculated elastic modulus was averaged, and standard deviation was determined. The elastic modulus of the sample indented by an AFM tip can be described by the Hertz model.62 The Hertz model gives the following relation between the indentation depth δ and the loading force F
and materials chemistry. Valuable insight was developed how dimensionality (2D vs 3D) HAP direct-write scaffolds, in terms of the architecture and subtle difference in surface roughness, stiffness, and chemical composition, can play a crucial role in governing the complex mechanism of modulation in osteoblast proliferation rate, osteoblast-to-osteocyte differentiation, timedependent downregulation in the expression of osteoblast markers, collagen type I and ALP, followed by the unregulated expression of MMP, osteocytic terminal differentiation markers (podoplanin, sclerostin), and dendritic extension formation. However, there remains a lot for further understanding of complex molecular mechanisms and detailed signaling pathways in the terminal differentiation of osteoblast accounting for alterations in the substrate so as to recapitulate the in vivo bone tissue microenvironment.
4. MATERIALS AND METHODS 4.1. Materials. HAP powder from Sigma-Aldrich (99.9% purity) (cat no. 12167-74-7) of reagent grade, having an average particle diameter of 4429 ± 382 nm, served as the colloidal phase in our ink. Poly(acrylic acid) (PAA) of SigmaAldrich (cat no. 9003-01-4) is an anionic polyelectrolyte used as a dispersant agent with deionized water to disperse HAP particles. Carboxymethyl cellulose of Merck (cat no. 217274) was used as a viscosifying agent. Calcium nitrate tetrahydrate (2 M) of Sigma-Aldrich (cat no. 13477-34-4) was used as a condensing agent. Ammonium hydroxide and nitric acid of Sigma-Aldrich (cat no. 320145 and 438073) were used to stabilize the pH of the solution. All chemicals used were of analytical grade and used without further purification. 4.2. HAP Ink Preparation. Prior to ink formation for direct writing, HAP powder was calcinated at 980 °C for 10 h in a muffle furnace, followed by ball-milling to break up particle agglomerates on a long-roll jar mill at 30 rpm for 30 h. Obtained HAP powders were sieved to get homogeneous particle size distribution. Sieved HAP particles dispersed in aqueous media containing PAA and water in the ratio (2:1), and the pH was adjusted to 9.0 with the addition of 5 M ammonium hydroxide. A 40 vol % of HAP powder was added in parts, followed by ultrasonication for 20 min to the solution for preparing colloidal suspension. At pH ∼9.0, PAA species were fully ionized, allowing them to impart the desired suspension stability.19 The suspension was stirred for 24 h and then centrifuged at 2000 rpm for 1 h to concentrate HAP. The supernatant was decanted, viscosifier (5 mg/mL) was added to increase the intrinsic viscosity, and 2 M calcium nitrate tetrahydrate was added to concentrate HAP to prepare ink. The resulted ink was suitable for printing. 4.3. Fabrication of HAP Scaffolds. Self-supportive 3D periodic HAP scaffolds were fabricated by extruding the respective ink through a stainless steel tip with a 260 μm diameter (Suzhou Lanbo Needle Co. Ltd, China), mounted onto the three-axis, computer-controlled robotic stage (Fiber Align, Aerotech Inc., Pittsburgh, USA) of DWA.58 The main advantage of direct-write technique is that the scaffold can be fabricated in one step using CAD design without using any additional mask, mold, or die.59 HAP ink was fed into the syringe and fixed in the barrel of DWA. The printing temperature was maintained at 25 °C, and the inks were extruded under the pneumatic pressure of 25−30 psi and at a writing speed of 0.3 mm/s, as controlled by a customized software.60 The overall set dimensions of scaffolds for printing were in the range of 25 mm2 in the x−y plane, center-to-center
i 2 yi E yz 2 zδ tan(α) F = jjj zzzjjj 2z k π {k 1 − ν {
(1)
where E is the elastic modulus, ν is the Poisson ratio, and α is the half-opening angle of the cylindrical cone for the AFM tip. A Poisson ratio of ≈0.3 was considered for HAP.61 A 25° was taken as the half-opening angle of the cone, corresponding to the manufacturer’s specifications. Equation 1 was used for obtaining a value for the elastic modulus E. F is calculated as F = xk
(2)
where x is the measured piezo movement and k is the spring constant. By using these equations, stiffness was measured 1512
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trophotometric analysis at 405 nm with n = 3 samples for each condition. Constructs cultured in osteogenic media were also analyzed for in vitro estimation of total collagen content with n = 3 samples for each condition. The total collagen content was estimated using a hydroxyproline assay. After enzymatic digestion of the constructs, the hydroxyproline content was quantified by treatment with p-dimethylaminobenzaldehyde (Sisco Research Laboratories Pvt. Ltd, cat. no. 044912) and chloramine-T (Central Drug House, cat. no. 027670) according to the manufacturer’s protocol. The hydroxyproline content in samples was calculated by referring to the standard curve generated in advance. Further, DNA content was estimated at 2nd, 4th, and 6th week. DNA was isolated from the constructs using a DNA extraction kit (Agilent) according to the manufacturer’s protocol.63 Purity of DNA and DNA concentration was estimated using a Nanodrop 2000C (Thermo Scientific, Wilmington, USA). 4.4.8. Quantitative Real Time Polymerase Chain Reaction. On 2nd, 4th, and 6th week, the total RNA was extracted from the engineered constructs using an RNeasy Mini Kit (Qiagen, cat. no. 74104). RNA concentration and the purity were determined using a Nanodrop spectrophotometer (Thermo Scientific). The extracted RNA was reverse-transcribed into cDNA using a first-strand cDNA synthesis kit (Thermo Scientific, cat. no. K1612). Quantitative real-time polymerase chain reaction (RT-PCR) was conducted using SYBR Green master mix (Quantitect, cat. no. 204074) and Rotor Gene Q thermocycler (Qiagen).64 The reactions were carried out in triplicates in 25 μL total volume containing 1 μL cDNA and 2.5 μL primer. The assay-on-demand (Qiagen) primers used were GAPDH (cat. no. QT00079247), ALP (cat. no. QT00211582), runt-related transcription factor (RUNX2) (cat. no. QT00020517), osteonectin (SPARC) (cat. no. QT00018620), osteopontin (cat. no. QT01008798), collagen type I (cat. no. QT00037793), BSP-2 (cat. no. QT00093709), osteocalcin (cat. no. QT00232771), osterix (cat. no. QT00213514), podoplanin (cat. no. QT01015084), β-catenin (cat. no. QT00077882), MMP-2 (cat. no. QT00088396), dentin matrix acidic phosphoprotein-1 (DMP-1) (cat. no. QT00022078), and SOST (cat. no. QT00219968). The analysis was carried out with the Rotor gene Q software, and the relative expression levels were calculated using the 2−(ΔΔc(t)) method with GAPDH as a reference gene. 4.4.9. Alizarin Red Staining and Immunofluorescence Analysis. On 4th week, constructs were harvested, rinsed in PBS, fixed in 10% formaldehyde, and stained with alizarin red dye (Sigma-Aldrich, USA, cat. no. A5533) for assessing the presence of mineralized nodules for osteogenic differentiation. Additionally, washed and fixed scaffolds were treated with 0.2% bovine serum albumin for blocking the nonspecific background staining. They were then processed for immunofluorescence using antibody against collagen type I (Millipore, cat. no. MAB3391) for osteogenesis and phalloidin (ECM Biosciences, cat. no. PF7551) for actin filaments and counterstained with 4′,6′-diamidino-2-phenylindole (DAPI) (Sigma-Aldrich, cat. no. 32670). Samples were visualized, and images were captured using Leica TCS SP5 (Leica Microsystems), an inverted confocal laser scanning microscope, equipped with an argon laser (457−514 nm), a diode laser (405 nm), and a DPSS laser (561 nm). The acquisition software was LAS AF software (Leica Microsystems).
three times at different locations and values were averaged to get a stiffness value for 2D and 3D HAP scaffolds. 4.4.3. Scanning Electron Microscope. The microstructure and morphological features of calcinated powder and sintered 2D and 3D HAP scaffolds were analyzed by SEM (ZEISS EVO 50). In order to get SEM images, scaffolds were washed with deionized water and ethanol. Gold coating was done by a gold sputter coater (EMITECH K550X, UK) for 1 min and 25 s with 25 mA electric current to form 15−20 nm thick coating. Different locations were scanned for the characterization of surface morphology and cell attachment on the scaffolds at varying magnifications. 4.4.4. X-ray Diffraction. XRD pattern of the calcinated powder and sintered 2D and 3D scaffolds was collected by using a Rigaku MiniFlex 600 benchtop X-ray diffractometer at step size 0.02° in the 2θ range 10−50° with target Cu Kα (λ = 1.54 nm). Target was set at 40 kV and 15 mA power level. Presence of different phases in powder, 2D, and 3D samples was identified by using JCPDS data files by using Pcpdfwin software maintained by International Centre for Diffraction Data (ICDD). 4.4.5. Attenuated Total Reflectance−Fourier Transform Infrared Spectroscopy. ATR−FTIR spectra of calcinated powder HAP and 2D and 3D sintered HAP scaffolds were recorded at room temperature by using an Alpha-P spectroscope (Bruker, USA). Transmission spectra were recorded in the wavelength range 400−4000 cm−1. Origin 8.5 software was used for correcting the baseline of each spectrum. 4.4.6. Cell Culture. Primary human osteoblasts (Lonza, cat. no. CC2538) were expanded in Petri dish at a density of 10 000 cells/cm2 in cell proliferation media consisting of Dulbecco’s modified Eagle’s medium (DMEM) (Himedia, cat. no. AL219A) supplemented with a 10% fetal bovine serum (Biological Industries, cat. no. 04-121-1A), HEPES buffer 1 M (Lonza, cat. no. 17-737E), 100 U/mL penicillin−streptomycin (Himedia, cat. no. A001A), 50 μg/mL gentamycin sulfate (Himedia, cat. no. A010), and 100 μg/mL amphotericin B (Himedia, cat. no. A011) and incubated at 37 °C and 5% CO2. Once the cells were subconfluent, the culture dish was washed with phosphate-buffered saline (PBS) (Himedia, cat. no. TL1006) and the osteoblasts were detached using 0.25% of trypsin EDTA (Himedia, cat. no. TCL007). All the scaffolds were incubated for 1 day in complete medium for preconditioning. The expanded primary human osteoblasts (P2) were seeded at a density of 1 × 105 cells/scaffold and cultured up to 6 weeks. The scaffolds were dipped into the cell proliferating media, followed by replacement with osteogenic differentiation media. Differentiation media contained DMEM with 10 nM dexamethasone (Sigma-Aldrich, cat. no. D2915), 0.01 M β-glycerol phosphate (Santa Cruz, cat. no. sc-203323), and 50 μg/mL ascorbic acid-2-phosphate (Sigma-Aldrich, cat. no. A8960). The medium was replenished every 3 days. The scaffolds were harvested at 2nd, 4th, and 6th week for analysis with n = 3 samples for each condition. 4.4.7. Biochemical Estimation and DNA Content. ALP biochemical assay kit (Bioassay Systems, cat. no. DALP-250) was used as per manufacturer’s protocol so as to determine the level of ALP released by osteoblast seeded on 2D and 3D HAP scaffolds at different time intervals. The ALP activity of osteoblasts was determined based on conversion of pnitrophenyl phosphate to p-nitrophenol, followed by spec1513
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4.4.10. Statistical Analysis. Each test was carried out in triplicates. Unless otherwise mentioned, data are presented as mean ± standard deviation. Statistical analyses were performed using a two-tailed Student’s t-test with p ≤ 0.05 and p ≤ 0.01 as the criteria for statistical significance.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. ORCID
Sourabh Ghosh: 0000-0002-1091-9614 Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This study was supported by funding from the Department of Biotechnology, Government of India (BT/PR8038/MED/32/ 303/2013 and BT/PR15451/MED/32/539/2016).
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DOI: 10.1021/acsomega.8b03272 ACS Omega 2019, 4, 1504−1515
