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J Dent Res 82(5): 372-376, 2003
© 2003 International and American Associations for Dental Research

RESEARCH REPORT
Biomaterials & Bioengineering

Implant Surface Roughness Affects Osteoblast Gene Expression

G.B. Schneider*, H. Perinpanayagam, M. Clegg, R. Zaharias, D. Seabold, J. Keller, and C. Stanford

N402, Dows Institute for Dental Research and the Department of Prosthodontics, University of Iowa College of Dentistry, Iowa City, IA 52242;

* corresponding author, galen-schneider{at}uiowa.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS & METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The transcription factor Cbfa1 regulates osteoblast differentiation and expression of genes necessary for the development of a mineralized phenotype. The purpose of this study was to determine if Cbfa1 and BSPII gene expression are influenced by implant surface microtopography. Osteoblasts were cultured on 600-grit (grooved) or sandblasted (roughened) cpTi implant discs. Mineralization was evaluated by Alizarin-Red-S staining. Real Time PCR was used for quantitative analysis of Cbfa1 and BSPII gene expression. Enhanced mineralization was seen in osteoblasts grown on roughened implant surfaces relative to tissue culture plastic. Real Time PCR showed significant (P < 0.05) increases in Cbfa1 gene expression in cells grown on roughened, as compared with grooved, implant surfaces. BSPII gene expression was also increased on rough surfaces in the UMR cells, but was reduced in the rat calvarial osteoblast cultures. These results suggest that osteoblast gene expression and mineralization are affected by roughened implant surface microtopographies during osseointegration of dental implants.

KEY WORDS: Cbfa1 • osseointegration • dental implants • osteoblasts • mineralization


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS & METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Many studies have been directed at attempts to understand the interaction of osteoblasts with various alloplastic biomaterials, such as titanium, that are often used to fabricate dental implants. Histological as well as biochemical data from these studies describe a variety of cellular responses to various implant surface conditions (Bowers et al., 1992; Schneider and Burridge, 1994; Boyan et al., 1996; Masuda et al., 1997).

During bone formation, the osteogenic marker Cbfa1 has been shown to be critical in the regulatory processes of bone differentiation (Ducy et al., 1997). Cbfa1 belongs to the Runt family of transcription factors (Xiao et al., 1998), and regulates osteoblast differentiation and expression of key osteoblast genes necessary for the development of a mineralized phenotype (Xiao et al., 1998; Ducy et al., 1999). However, the effects of different implant surface microtopographies on gene expression of key osteogenic factors are not fully understood.

The hypothesis of the current study was that roughened implant surface microtopographies differentially affect Cbfa1 and BSPII gene expression, as well as mineralization. We utilized a previously characterized, rapidly mineralizing UMR-106-01 BSP osteoblast model (Stanford et al., 1995), and a non-transformed primary rat calvarial osteoblast (RCOB) model for comparison (Bowers et al., 1992). In the current study, rapidly mineralizing UMR-BSP osteoblasts were grown on either rough or grooved implant surfaces. Enhanced mineralization, as well as increased gene expression of Cbfa1 and BSPII, was noted on rougher surfaces as compared with grooved, relative to tissue culture plastic. However, in the RCOB cultures, less BSPII gene expression was seen on rough surfaces relative to grooved. These results suggest that osteoblast gene expression and subsequent mineralization are affected by roughened implant surface microtopographies during the osseointegration of dental implants.


   MATERIALS & METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS & METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture and Titanium Implant Specimens
UMR-106-01-BSP and primary rat calvarial osteoblast cell cultures were grown, passaged, and maintained as previously described (Bowers et al., 1992; Stanford et al., 1995). A non-mineralizing subclone of the parental UMR-BSP cell line, UMR-UI cells, was used for the negative mineralization control (Schneider et al., 2001). Commercially pure titanium (cpTi) discs with dimensions of 1.25 cm diameter x 4 mm height were used. The disc surfaces were polished to either a rough (sandblasted) or grooved (600 grit) finish as previously described (Bowers et al., 1992).

Alizarin Red S Staining for Mineralization
UMR osteoblasts were plated in triplicate at a density of 50,000 cells/10-µL droplet cultures onto prepared titanium discs or onto tissue culture plastic and placed into a 37-degree incubator. After 1 hr of attachment, 1.0 mL of medium was added to each culture. Following an incubation period of 48 hrs, ß-glycerophosphate (5 mM final concentration) was added, and the cultures were incubated for an additional 24 hrs. The medium was then removed, and the culture was gently rinsed with phosphate-buffered saline (PBS) and fixed for 10 min in 10% formalin. After fixation, the cultures were rinsed with Nanopure water (no calcium ion) and stained for calcium with 2% Alizarin Red-S (AR-S) for 10 min at room temperature. Wells were washed 4x with Nanopure water and evaluated for intensity of AR-S staining. Triplicate cultures were scanned for quantitation by means of the NIH Scion imaging analysis program. Each culture was then scanned 3x and averaged. The averaged scan value of the non-mineralizing control (UI cells) was subtracted from all BSP scan values. The experimental BSP cultures were expressed as a percentage of the BSP on the rough-surface group. Statistical analysis (N = 3) was performed by a one-way analysis of variance (ANOVA) with Tukey’s Multiple Comparison Test to a significance level of P < 0.05.

Cell Proliferation and Attachment Assay
Microdots of UI cells and BSP cells were plated in triplicate (50,000 cells/10-µL dot) on tissue culture plastic (TCP) and on cpTi discs polished to either a rough (sandblasted) or grooved (600 grit) finish. Cultures were allowed to attach for 1 hr before being flooded with 1 mL of medium. At 48 hrs, 5 mM ß-glycerophosphate was added to all cultures. At 72 hrs, the medium was removed from the wells, unattached cells were quantitated with the model ZM Coulter counter, and triplicate cultures were averaged. Statistical analysis (N = 5) was performed by one-way analysis of variance (ANOVA) with Tukey’s Multiple Comparison Test to a significance level of P < 0.05.

Real Time RT-PCR
Real Time PCR primers and probes were designed (Fig. 1Go) with Primer Express software (Perkin Elmer, Boston, MA, USA) from the known rat sequence that corresponds to exons 1 and 2 (Isoform 1), and 3 and 4 (Isoform 3, Runt-domain) of the Cbfa1 gene (Xiao et al., 1998), as well as exons 6 and 7 of BSPII (Oldberg et al., 1988). The probe was designed to overlie the exon junction, so that it was unlikely to hybridize to genomic DNA that may have been present. This ensured that if genomic DNA contaminants were present, their PCR amplification was non-contributory to the Real Time analysis. We used the TaqMan Ribosomal RNA Control Reagents Kit (Perkin Elmer) to detect 18s ribosomal RNA as an endogenous control.



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  		Figure 1. Design of Real Time primers for Cbfa1 and bone sialoprotein.

 
RNA extracts were analyzed in triplicate by quantitative Real Time Multiplex RT-PCR by means of a TaqMan Gold RT-PCR Kit (Perkin Elmer). The RNA was reverse-transcribed into cDNA at 48°C for 30 min. Then, the Cbfa1 or BSPII target and the endogenous rRNA control were amplified by multiplex PCR with thermal cycling parameters of 50°C for 2 min, 95°C for 10 min, 40 cycles of 95°C for 15 min, and 60°C for 1 min. The Real Time PCR reactions were performed in 96-well Optical Reaction Plates (Perkin Elmer) in an ABI Prism 7700 Sequence Detection System (Perkin Elmer). PCR product was separated on an agarose gel (4%), stained with ethidium bromide, and visualized on a Transilluminator. The band was excised and purified with the QIAquick Gel Extraction Kit (Qiagen, Valencia, CA, USA), sequenced in an Applied Biosystems Model 377 DNA sequencer at the University of Iowa DNA facility, and confirmed, by Blast Search, to be Cbfa1 or BSPII. Steady-state mRNA levels of each isoform were normalized to 18s rRNA and calculated by the Comparative Method of Relative Quantitation in Multiplex Reactions (Perkin Elmer). Statistical analysis (N = 9, 3 groups repeated 3 times) of {triangleup}{triangleup}Ct was performed by a one-way analysis of variance (ANOVA) with Tukey’s Multiple Comparison Test or a paired one-tailed t test to a confidence level of P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS & METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Osteoblast Mineralization is Greater on Roughened Implant Surfaces
Titanium discs were polished to either a rough (Fig. 2AGo) or grooved (Fig. 2BGo) finish. Triplicate UMR-BSP cultures were grown for 72 hrs on these prepared implant surfaces. A non-mineralizing subclone, UMR-UI, cultured on tissue culture plastic, was used as a negative control for mineralization comparison (Fig. 3AGo). Alizarin Red-S stain for mineralization showed that osteoblasts cultured on rough implant surfaces had increased mineralization, as shown by a more intense red staining (Fig. 3AGo). When quantitated by an NIH image analysis program, a significant (P < 0.05) increase in mineral of nearly 50% was noted in osteoblasts cultured on rough as compared with grooved implant surfaces (Fig. 3AGo).



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  		Figure 2. SEM of the sandblasted rough (A) and 600-grit grooved (B) surface structures of the prepared titanium discs. Magnification, 1000x; bar = 10 µm.

 




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  		Figure 3. Osteoblast cultures mineralize significantly more (P < 0.05) on rough as compared with grooved implant surfaces, as demonstrated by Alizarin Red S staining (A). Reduction in mineralization was not due to loss of cell attachment (B), or differences in rates of cell proliferation (C). n = 5/group; mean ± SD.

 
To confirm that the difference in mineralization was not due to cell loss or differences in rates of cell proliferation, we performed cell attachment and proliferation assays at 72 hrs. No significant difference in cell attachment was evident between osteoblasts cultured on rough or grooved implant surfaces, thus demonstrating that the difference in mineralization on grooved vs. rough surfaces was not due to the presence of fewer cells (Fig. 3BGo). We did not find significant differences in the proliferative rates of osteoblasts cultured on the surfaces tested (Fig. 3CGo).

Cbfa1 and BSPII Gene Expression are Affected by Implant Surface Microtopographies
Differences in levels of mineralization were noted in osteoblasts cultured on rough vs. grooved implant surfaces. We used Real Time RT-PCR strategies to determine if differences in the levels of Cbfa1 and BSPII gene expression would also differ (Fig. 4Go). Quantitative Real Time PCR analysis detected Cbfa1 and BSPII in the mineralizing osteoblast cultures grown on both rough and grooved implant surfaces, relative to tissue culture plastic. The steady-state mRNA levels of Cbfa1 were significantly (P < 0.03) greater, nearly four-fold, in the UMR osteoblast cultures grown on the roughened implant surface as compared with grooved (Fig. 4AGo). This result was also found in the non-transformed RCOB cultures, where, again, a significant (P < 0.001), three-fold increase in Cbfa1 gene expression was noted on the rough surface (Fig. 4BGo). In addition, the expression of the extracellular matrix protein specific to mineralizing tissues, BSPII, whose gene expression is thought to be regulated by the Cbfa1 transcription factor (Ducy et al., 1997), was slightly increased (P < 0.01) in UMR osteoblasts grown on the roughened surface as well (Fig. 4CGo). However, when analyzed in the RCOB cultures, there was six-fold less (P < 0.001) BSPII gene expression on rough surface microtopographies relative to grooved (Fig. 4DGo).



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  		Figure 4. Cbfa1 gene expression is significantly (P < 0.05) increased in UMR (A) and RCOB (B) osteoblast culture models grown on roughened implant surface microtopographies as demonstrated with Real Time PCR analysis. BSPII gene expression increased in UMR cultures (C), but was reduced in RCOB cells (D) cultured on rough as compared with grooved implant surfaces. n = 9/group; mean ± SD.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS & METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Osteoblast differentiation, ECM formation, and subsequent mineralization are needed for bone formation associated with osteogenesis and subsequent osseointegration. The transcription factor Cbfa1 regulates this development (Ducy, 2000), which, if disrupted, leads to the absence of osteoblasts and lack of bone development (Komori et al., 1997). Cbfa1 binds to the OSE2 promoter region of major osteoblast genes such as osteocalcin, bone sialoprotein, osteopontin, and type I collagen and regulates their expression (Ducy et al., 1997, 1999).

Osteoblast differentiation and responses during osseointegration vary and are affected by the implant surface microtopography, associated extracellular matrix proteins, and their respective integrin receptors (Schneider and Burridge, 1994; Okamoto et al., 1998; Cooper et al., 1999; Sykaras et al., 2000; Schneider et al., 2001). A direct association of osteoblasts with their associated bone matrices defines osseointegration with the implant surface in the absence of soft or fibrous tissue (Albrektsson et al., 1981). Many studies examining cell adhesion and morphology, DNA synthesis, integrin and extracellular matrix expression, and enzyme activity have been done to elucidate osteoblastic response to titanium alloys (Puleo et al., 1991; Vrouwenvelder et al., 1993; Schneider and Burridge, 1994; Gronowicz and McCarthy, 1996; Sinha and Tuan, 1996). However, many of the molecular and genotypic events taking place at the osteoblast cell level during osseointegration are still largely unknown.

The purpose of our study was to begin to address these molecular events with respect to Cbfa1 and BSPII gene expression in relation to differences in mineralization associated with roughened implant surface microtopographies. Data from this study provide novel cellular and molecular insight as to the effects of implant surface microtopographies, in association with the surrounding matrix, on molecular signaling mechanisms that regulate osteoblast mineralization during osseointegration.

Following the preparation of two different implant surface conditions, rough and grooved, we utilized the UMR-BSP-106-01 osteoblast cell model (Stanford et al., 1995; Schneider et al., 2001) and the non-transformed primary RCOB model to assess differences in levels of mineralization when osteoblasts were cultured onto these prepared implant surfaces. Osteoblasts grown on roughened surfaces, as previously reported (Bowers et al., 1992), revealed more mineralization as compared with those on grooved surfaces.

As a direct result of these differences in mineralization, and because of the known function of Cbfa1 in the regulation of osteoblast differentiation and subsequent mineralization, we wanted to determine if differences in levels of Cbfa1 expression would also be seen under in vitro conditions simulating osseointegration. To analyze and quantify the results, we used Real Time RT-PCR strategies that can determine absolute mRNA copy number (Aarskog and Vedeler, 2000; Bustin, 2000). Cbfa1 gene expression was enhanced in the UMR and rat calvarial osteoblasts grown on the rough implant surfaces relative to grooved. Analysis of the suggested Cbfa1 transcriptionally regulated BSPII gene (Ducy et al., 1997) revealed slight enhancement of expression in UMR osteoblasts, but reduced expression in RCOBs on the rough surfaces relative to grooved. However, expression of both genes was higher on both types of rough implant surfaces relative to tissue culture plastic. Differences in BSPII gene expression between the UMR and rat calvarial osteoblasts could be for several reasons. The expression of Cbfa1 might be higher in the RCOBs cultured on the rough surface microtopographies, as compared with the grooved, but the permissive activity of this Cbfa1 transcription factor might be impeded as a result of an unknown effect of the rough, but not the grooved, implant surface microtopography on this intracellular signaling cascade. This impediment in Cbfa1 function, but not expression, may lead to a reduction of BSPII gene expression as seen in the RCOB cultures on the rough relative to the grooved surfaces. The impediment of Cbfa1 transcription factor with a BSPII gene could also be associated with interference of Cbfa1 with the recently described osterix gene (Nakashima et al., 2002).

Additionally, on the grooved surface, Cbfa1 was expressed at a level similar to that of BSPII gene expression on the same surface in the RCOBs, where, as in the UMR model, a slight increase in BSPII gene expression relative to Cbfa1 was noted. The modest increase in BSPII gene expression relative to Cbfa1 in the UMR model, vs. the reduction of BSPII relative to Cbfa1 on rough surfaces in the RCOB cells, may be attributed to the transformed state of the UMR model, where overexpression of BSPII has been reported, and thus may be an artifact of the culture model (Stanford et al., 1995). Thus, the UMR cells may be able to override the lack of Cbfa1 permissive activity seen in the RCOB model on rough microtopographies by the ability of the UMR model to overexpress that BSPII gene as a function of its transformed state. Differences in gene expression on rough and grooved microtopographies in both the UMR and RCOB cells were not attributed to variations in rates of cell proliferation or number, since no significant differences in these variables were noted.

We show here that different implant surface microtopographies (rough vs. grooved) can alter the expression of key osteogenic regulatory genes such as Cbfa1 and BSPII. This suggests that the interaction of the osteoblasts with the extracellular matrix components on the different implant surface microtopographies can influence gene expression. Perhaps this occurs as a result of differences in cell adhesion and shape, as a result of integrin-mediated adhesion and regulation of downstream signaling cascades as previously reported (Schneider et al., 2001). It could also be a result of extracellular matrix spatial and temporal expression profile changes that would be under the control of the transcription factor Cbfa1, such as bone sialoprotein (BSPII) (Ducy et al., 1997; Schneider et al., 1999).

Thus, roughened implant surface microtopographies may contribute to the regulation of osteoblast differentiation by influencing the level of gene expression of key osteogenic factors. A better understanding of these molecular processes will lead to the development of more advanced therapeutic prosthetic interventions associated with dental implant therapy and tissue-engineering biological applications.


   ACKNOWLEDGMENTS
 
The authors thank Kevin Knudtson and the University of Iowa DNA Facility for guidance with the Real Time PCR analysis. This study was supported by the University of Iowa College of Dentistry Research Foundation (GS), the NIH/NIDCR grant P3010126-09 (GS), the NIH/NIDCR grant P60DE13076 (GS), and the Roy J. Carver Charitable Trust (CS).

Received January 15, 2002; Last revision February 11, 2003; Accepted February 11, 2003


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