Research Article

J Med Discov (2018); 3(3):jmd18024; DOI:10.24262/jmd.3.3.18024; 
Received May 6th, 2018, Revised May 27th, 2018, Accepted June 10th, 2018, Published July 3rd, 2018.

Folic Acid-Modulated Growth of Dental Pulp Stem Cells (DPSCs)

Kristi Agari¹, Weiye Lin¹, Karl Kingsley¹

¹University of Nevada, Las Vegas – School of Dental Medicine, 1001 Shadow Lane, Las Vegas, NV 89106

* Correspondence: Karl Kingsley, Professor of Biomedical Sciences and Director of Student Research, University of Nevada, Las Vegas – School of Dental Medicine 1001 Shadow Lane, B313, Las Vegas, Nevada 89106. Phone:(702) 774-2623. Fax:(702) 774-2721. E-mail: Karl.Kingsley@unlv.edu.

Folates are associated with a wide variety of human health benefits, including reduced risk for stroke, heart failure, and coronary heart disease as well as chemo-preventive and anti-tumor actions of folates. Recent evidence has suggested  folic acid (FA) may facilitate growth and protect cellular viability in mesenchymal stem cells, although no studies have yet examined this potential in dental pulp-derived stem cells (DPSCs). Using multiple DPSC isolates, FA was administered at supraphysiologic levels corresponding with bioavailable concentration following FA supplementation [0-400 ug/mL]. Cellular growth and proliferation, as well as viability were assessed. In addition, key enzymes involved in FA metabolism were also assessed using RT-PCR. These data reveal that FA administration is sufficient to induce significant increases in cellular proliferation at both low and high FA concentrations [100, 400 ug/mL]. In addition, dose-dependent increases in cellular viability were also observed. These changes correlated with an increase in mRNA expression of dihydrofolate reductase (DHFR) and thymidylate synthetase (TS), which are responsible for distinct intracellular pathways of folate metabolism. These observations may provide novel information regarding the ability of FA to stimulate growth and increase viability in DPSC, which may be critical for future studies of regeneration and bioengineering for clinical and translational therapies. These data may provide novel information to facilitate positive homeostatic microenvironments useful to develop and differentiate DPSCs for therapeutic and clinical treatments.

Keywords: Dental Pulp Stem Cells (DPSC), Folic Acid (FA), growth, viability

Folates are associated with a wide variety of human health benefits, including improved insulin resistance and glycemic control, improved cognitive functions and reduced symptoms associated with Alzheimer-related dementia, as well prevention of neural tube defects in pregnancy [1-4]. Other studies have focused on the cardiovascular disease prevention aspects of folate intake and dietary supplementation, demonstrating reduced risk for stroke, heart failure, and coronary heart disease [4,5]. Moreover, evidence continues to accumulate regarding the chemo-preventive and anti-tumor actions of folates, with substantial evidence supporting these effects to prevent and ameliorate colorectal cancers [6,7].

Despite the breadth and depth of research regarding the positive outcomes and benefits of folate intake and dietary folate supplementation, lack of dietary folate continues to be a risk factor for many health problems, including congenital defects, developmental disorders, and many human cancers [8-12]. Although much of these efforts have focused on colorectal cancer, risk for many other cancers including breast cancers and leukemia may also be linked to folate deficiency [13-15]. In fact, some evidence now suggests that folate deficiency may affect the specific response of cancer stem cells thereby increasing the risk of chemotherapeutic resistance and metastatic potential [16-18].

These observations may provide support for observations of mesenchymal stem cells, which may also respond differentially to folate supplementation and deprivation [19-23]. Although these studies have evaluated the potential effects of folic acid (FA) to facilitate growth and protect cellular viability in mesenchymal stem cells, to date no studies have examined this potential in dental pulp-derived stem cells (DPSCs). Based upon the paucity of evidence, the goal of this study was to assess the effects of FA supplementation on human DPSCs.

Human subjects
Dental pulp stem cells were obtained from patients at the University of Nevada, Las Vegas – School of Dental Medicine (UNLV-SDM) dental clinic. This study was reviewed and approved by the Institutional Review Board (IRB) in the Office for the Protection of Research Subjects (OPRS). Approval for the original study “Isolated of Non-Embryonic Stem Cell s from Dental Pulp” was obtained in February 2010 (OPRS 0907-3148).

In brief, subjects were recruited from clinic patients of record at UNLV-SDM that were previously scheduled for tooth extraction of one or more healthy, vital intact teeth – mainly for Orthodontic treatment. Prior to participation, all potential subjects provided Informed Consent. Any patients that were found to have injured or compromised (fractured, diseased) teeth and those patients that refused to participate were excluded.

Cell culture and DPSC identification
Following extraction, each tooth was sectioned to expose the dental pulp that was subsequently extracted and deposited into a sterile microcentrifuge tube containing sterile saline solution for transfer to the laboratory for culture using the direct outgrowth method, as previously described [24-26]. Subsequent culture was facilitated using RPMI-1640 culture media with 2 mM L-Glutamine containing 4.5 g/L glucose, 1.5 g/L sodium bicarbonate, 10 mM HEPES, 1.0 mM sodium Pyruvate, and 1% Penicillin-Streptomycin (10,000 unit/mL), as previously described [24-29]. Analysis of the resulting DPSC isolates after 10 passages demonstrated each patient isolate exhibited a distinct proliferation rate or doubling time (DT), which were characterized as rapid or rDT (less than three days), slow or sDT (greater than one week, usually 8 – 10 days), and a few with intermediate or iDT [25,29,30]. Most of the isolates exhibited rDT (n=25), while fewer exhibited iDT (n=2) or sDT (n=3) [29,30]. Cultures with greater than 1.0 x 10⁶ cells/mL were considered confluent and were passaged or split 1:2. All DPSC isolates were screened for the expression of Sox-2, NANOG, and Oct4 – biomarkers for DPSC and MSC [24,25,27-30].

Cell proliferation and viability
Cells were assessed for viability using the Trypan Blue method, which involves mixing an aliquot of non-adherent DPSC for analysis in a BioRad TC20 automated cell counter and hemocytometer using the protocol recommended by the manufacturer, which provides information regarding cell number, relative abundance (confluence) and percent viability [31,32]. Cultures with greater than 1.0 x 10⁶ cells/mL were considered confluent and were passaged or split 1:2. All cultures utilized for each DPSC isolate were P10 or greater.

Folic acid
Folic acid (folate) molecular weight (MW) 441.40 was obtained from Fisher Scientific (Fair Lawn, NJ) and suspended in the appropriate cell culture complete media at concentrations between 100 – 400 ug/mL that approximated the supraphysiologic range 0.23 mM  to 0.91 mM, which is observed under conditions of folate supplementation [33,34]. Cells were plated in 96-well assay plates at 1.2 x 10⁵ cells/mL with eight (n=8) replicates for each condition. Three (n=3) separate, independent experiments were performed for each isolate, folate concentration and experimental condition, yielding a total of n=24 data points (8 replicates x 3 experiments) for each experimental condition. Cell number, relative confluence and percent viability were assessed to determine any changes to proliferation rate (doubling time, DT) and viability.

100 ug/mL = 100 mg/L = 0.1 g FA/441.4 MW = 0.0002265 or 0.23 mM

400 ug/mL = 400 mg/L = 0.4 g FA/441.4 MW = 0.0009062 or 0.91 mM

Statistics
Basic descriptive statistics were provided for cellular viability and proliferation rate (doubling time) and were compiled. Changes in cellular viability were measured using two-tailed Student’s t-tests with significance set at alpha level of a=0.05, which provide robust analysis for samples with even moderate samples sizes (n~20) [35,36].

RNA isolation and PCR
RNA was extracted from negative controls and experimental cells at each folic acid concentration (100, 400 ug/mL) using the ABgene total RNA isolation reagent (Surrey, UK) using the protocol recommended by the manufacturer, as previously described [24,25,27,28,33]. The quality (purity) and quantity of RNA was determined using spectrophotometric analysis at 260 and 280 nm. Ratio of the absorbance readings (A260, A280) will provide estimated purity, which should be approximately 1.70 or higher. RNA concentration can then be determined using the formula:

A260 (absorbance) x 40 (RNA standard) x dilution factor (50) = RNA yield per mL

RT-PCR was then used to screen the RNA from negative control and experimental cells to determine if any changes to mRNA expression were induced by the addition of folic acid. Equal amounts of RNA (1 ng) from each isolate used to screen for the positive control internal cell standard, which was glyceraldehyde-phosphate dehydrogenase or GAPDH (glycolytic enzyme). Each sample was then subsequently screened using primers specific for dihydrofolate reductase (DHFR), which is required to convert folate for the de novo synthesis of purines and thymidylate synthetase (TS) – the sole de novo pathway to produce dTMP in DPSC [34,37,38].

GAPDH forward primer, GGTCGGAGTCAACGGATT; 18 nt, 56% GC, Tm = 64C
GAPDH reverse primer, ATCGCCCCACTTGATTTTG; 19 nt, 47% GC, Tm = 63C
Annealing temperature = 64C
Optimal Tm (GAPDH-F: GAPDH-R) Lower Tm – 5C = 58C

b-actin forward primer, AGGGCAGTGATCTCCTTCTGCATCCT; 26 nt, 54% GC, Tm = 74C
b-actin reverse primer, CCACACTGTGCCCATCTACGAGGGGT; 26 nt, 62% GC, Tm = 77C
Annealing temperature = 72C
Optimal Tm (b-actin -F: b-actin -R) Lower Tm – 5C = 69C

DHFR forward primer, GGCGGCAAGCTTATGGTTGGTTCGCTAAACTGC; 33 nt, 55% GC, Tm = 78C
DHFR reverse primer, GCCGCCGGATCCATCATTCTTCTCATACTTC; 31 nt, 52% GC, Tm = 74C
Annealing temperature = 72C
Optimal Tm (DHFR-F: DHFR-R) Lower Tm – 5C = 69C

TS forward primer, GCCTCGGTGTGCCTTTCA; 18 nt, 61% GC, Tm = 69C
TS reverse primer, CCCGTGATGTGCGCAAT; 17 nt, 59% GC, Tm = 66C
Annealing temperature = 67C
Optimal Tm (TS-F: TS-R) Lower Tm – 5C = 61C

Four DPSC isolates were selected for inclusion in this initial pilot study (Table 1). Due to the higher prevalence of DPSC isolates with rDT (n=25/30 or 83.3%), two of the isolates chosen for inclusion exhibited rDT of approximately two days (dpsc-17322, dpsc-11836). Due to the lower prevalence of DPSC isolates with slower doubling times, the remaining two isolates chosen for inclusion each exhibited iDT of approximately six days (dpsc-5653) and sDT of nearly nine days (dpsc-7089). Baseline viability following cryopreservation, thawing and cell culture varied between 58% and 63%.

Table 1. Dental Pulp Stem Cell isolates

To evaluate the effects of FA the four DPSC isolates were plated in 96-well plates with or without the addition of FA at low [100 ug/mL] and high [400 ug/mL] concentrations and proliferation was observed over four days (Figure 1). The addition of FA at both low and high concentrations was sufficient to induce changes to proliferation in all four DPSC isolates. More specifically, the low concentration of FA increased growth in the rDT DPSC isolates dpsc-11836 (Fig. 1A) and dpsc17322 (Fig. 1B) between 1.56- and 1.70-fold, which was statistically significant, p<0.05. This same concentration also increased growth in the iDT and sDT DPSC isolates dpsc-5653 (Fig. 1C) and dpsc-7089 (Fig. 1D) between 1.40- and 1.61-fold, which was also statistically significant, p<0.05.

The additional of FA at the higher concentration (400 ug/mL) increased growth in the rDT DPSC isolates dpsc-11836 (Fig. 1A) and dpsc17322 (Fig. 1B) between 2.61- and 2.63-fold, which was statistically significant, p<0.01. This same concentration also increased growth in the iDT and sDT DPSC isolates dpsc-5653 (Fig. 1C) and dpsc-7089 (Fig. 1D) between 2.81- and 2.23-fold, which was also statistically significant, p<0.01.

Figure 1. FA-mediated changes in proliferation of DPSC isolates. 
A) Low (100 ug/mL) and high (400 ug/mL) concentrations of FA induced significant increases in rDT dpsc-11836, 1.56- and 2.61-fold. 
B) Low and high FA concentrations induced similar increases in rDT dpsc-17322, 1.70- and 2.63-fold. 
C) The addition of FA also stimulated iDT growth in dpsc-5653 by 1.40- and 2.81-fold. 
D) FA induced similar changes to sDT dpsc-7089, 1.61- and 2.23-fold. All concentrations evaluated induced statistically significant changes compared with negative controls, p<0.05.

To assess whether these changes to growth and proliferation observed in all DPSC isolates in response to FA administration also correlated with changes in cell survival, viability was also assessed at each time point and for each concentration (Figure 2). In brief, FA at the low concentration of 100 ug/mL induced changes in DPSC viability, increasing on average between 10% – 21%, which was statistically significant, p<0.05. In addition, FA at the higher concentration of 400 ug/mL induced greater increases in DPSC viability between 26% – 31%, which was also statistically significant, p<0.01.

Figure 1. FA-mediated changes in proliferation of DPSC isolates. A) Low (100 ug/mL) and high (400 ug/mL) concentrations of FA induced significant increases in rDT dpsc-11836, 1.56- and 2.61-fold. B) Low and high FA concentrations induced similar increases in rDT dpsc-17322, 1.70- and 2.63-fold. C) The addition of FA also stimulated iDT growth in dpsc-5653 by 1.40- and 2.81-fold. D) FA induced similar changes to sDT dpsc-7089, 1.61- and 2.23-fold. All concentrations evaluated induced statistically significant changes compared with negative controls, p<0.05.

Figure 2. FA-induced changes to DPSC viability. 
Low concentrations of FA [100 ug/mL] increased DPSC viability between 10% – 21% compared with baseline and negative controls. 
High concentrations of FA [400 ug/mL] increased DPSC viability between 26% - 31% compared with baseline and negative controls. These changes were statistically significant, p<0.05.

To determine how FA may be inducing these changes in cellular growth and viability, the two main metabolism enzymes responsible for folate metabolism were assessed using RT-PCR, dihydrofolate reductase (DHFR) – which is required to covert folate for the de novo synthesis of purines, and thymidylate synthetase (TS) – the sole de novo pathway to produce dTMP in DPSC (Figure 3). In brief, these results demonstrated that DHFR is produced in all four DPSC isolates, but that mRNA expression of DHFR is strongly enhanced following FA supplementation at either low [100 ug/mL] or high [400 ug/mL] FA concentrations (Fig. 3A). Similar results were observed for TS mRNA, which is also produced in all four DPSC isolates, but is strongly enhanced following FA supplementation at both low and high FA concentrations (Fig. 3B). Internal controls for structural (actin) and enzymatic mRNA included for reference standards to normalize DHFR and TS expression (Fig. 3C).

Although many studies have evaluated the potential effects of FA to facilitate growth and protect cellular viability in mesenchymal stem cells [19-23], to date no studies had examined this potential in dental pulp-derived stem cells (DPSCs). Based upon the paucity of evidence, the goal of this study was to assess the effects of FA supplementation on human DPSCs. These results clearly demonstrated that FA is sufficient to induce significant increases in cellular growth and proliferation, which are also correlated with increases in cellular viability.  Moreover, these increases appear to be, dose-dependent with higher concentrations of FA correlated with larger increases in growth and higher levels of viability.

In addition, this study also provides some of the first mechanistic evidence to demonstrate that FA administration is also inducing changes to enzymes directly responsible for the metabolism of FA within the cell.  Without these data, it might have been possible to suggest that FA could induce these changes through interactions with other intracellular pathways. However, the observed changes in mRNA for separate pathways of FA metabolism clearly suggest that the increased availability of FA may be directly linked with increased bioavailability and metabolism, which in turn may be directly altering cellular growth and viability.

Figure 3. FA-induced changes to folate metabolism enzymes. 
A) FA administration induced significant increases in mRNA expression of dihydrofolate reductase (DHFR), with an average increase in signal band intensity of approximately 3.24-fold at 100 ug/mL and 7.68-fold at 400 ug/mL. 
B) FA administration also induced significant increases in mRNA expression of thymidylate synthetase (TS), with an average increase in signal band intensity of 1.76-fold at 100 ug/mL and 2.71-fold at 400 ug/mL.

Despite these observations, there are some limitations of this study which should be considered.  First, this is an in vitro study of previously collected DPSC isolates [24,25].  These same observations may not be applicable in vivo and therefore may not represent a true representation of how DPSC may react to FA supplementation in the body. Furthermore, this study sample size was limited due to funding considerations – which suggests that more robust analyses of larger numbers of DPSC isolates will be needed in order to confirm and corroborate these findings.

New research has suggested DPSC may be used to repair or replace injured and aging tissues, such as nerves and bone – in addition to dental tissues and structures [39-42]. Any method proven to stimulate growth or increase viability may be critical for future studies regarding the regeneration and bioengineering of DPSC for clinical and translational therapies. These data may provide novel information to facilitate positive homeostatic microenvironments to develop and differentiate DPSCs for therapeutic and clinical treatments.

The authors declare that they have no competing interests.

The authors would like to acknowledge the UNLV-SDM Office of Research and the UNLV Graduate and Professional Student Association (GPSA) for funding of this study. KA acknowledges funding support by the National Institute Of Dental & Craniofacial Research of the National Institutes of Health under Award Number R13DE024621. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

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