University of Connecticut OpenCommons@UConn

SoDM Masters Theses School of Dental Medicine

2008 Characterization of Chemical and Biological Properties of Purified orP phyromonas gingivalis Free Dihydroceramides Andrew Robert Chapokas University of Connecticut Health Center

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Recommended Citation Chapokas, Andrew Robert, "Characterization of Chemical and Biological Properties of Purified Free Dihydroceramides" (2008). SoDM Masters Theses. 161. https://opencommons.uconn.edu/sodm_masters/161

Characterization of Chemical and Biological Properties of Purified Porphyromonas

gingivalis Free Dihydroceramides

Andrew Robert Chapokas

D.M.D., University of Pennsylvania, 2005

A Thesis

Submitted in Partial Fulfillment of the

Requirements for the Degree of

Masters of Dental Science

at the

University of Connecticut

2008

APPROVAL PAGE

Masters of Dental Science Thesis

Purification and Characterization of Chemical and Biological Properties of

Porphyromonas gingivalis Free Ceramides

Presented by

Andrew Robert Chapokas, D.M.D.

Major Advisor Frank Nichols, D.M.D., Ph.D.

Associate Advisor John Dean III, D.M.D., Ph.D.

Associate Advisor Qiang Zhu, D.D.S., Ph.D.

University of Connecticut

2008

ii

To my fiancé, Monica, for providing me with love and support,

And

To my family, Robert, Catherine, and Ashley who have nourished and encouraged me throughout my educational journey.

Without their help, this work would not have been possible.

iii

ACKNOWLEDGEMENTS

I would like to extend great thanks to my friend and major advisor, Dr. Frank Nichols, for the education he has provided me both in research and . Also, I thank my associate advisors, Drs. Robert Clark, John Dean, and Qiang Zhu whose time and additional recommendations are deeply appreciated.

In addition, I would like to thank Amber Onorato, Michael Smith, Martha Morton, and Marvin Thompson from the University of Connecticut, Storrs graduate chemistry department for kindly providing me with synthetic free dihydroceramide and its NMR spectra, and access to their electrospray mass spectrometer. I would also like to extend thanks to my co-residents and faculty in both periodontology and prosthodontics for their support and encouragement.

iv TABLE OF CONTENTS

Acknowledgements……………………………………………...... ………..iv

Abstract…………………………………………………………………... ..vi

Introduction…………………………………………………….…………. 1

Objective………………………………………………………….………...12

Materials and Methods………………………………………………...…… 13

Results……………………….…………………………………………….. 21

Discussion……………..……………………………………………….….. 24

Conclusion……………………………………………………………….… 32

Figures………………………………………………………………………33

References…………………………………………………………...... 47

v

ABSTRACT

The aim of the present investigation is to purify, characterize, and determine the chemical and biological properties of Porphyromonas gingivalis free dihydroceramides (fDHC). Effects were tested by examination of human gingival fibroblast morphologic changes in addition to their synthesis of prostaglandin E2

(PGE2) and prostaglandin F2α (PGF2α) produced in vitro. Free dihydroceramides as

well as phosphoglycerol dihydroceramide (PGDHC), and de-esterified phosphoglycerol dihydroceramide (PGDHC-C15:0) were obtained by total lipid

extraction of P. gingivalis followed by high performance liquid chromatography

(HPLC). Confirmation and structural analysis was obtained by electrospray mass

spectrometry (ES-/+MS) and nuclear magnetic resonance spectroscopy (NMR). A synthetic free dihydroceramide (A) was obtained, and its purification and structural

analysis was similar. Ethanol solvent served as a negative control, and PGDHC

served as a positive control. HGF were obtained from tissue samples of two

periodontally healthy volunteers during clinical procedures, and

cell cultures were introduced to 35 mm wells previously coated with ethanol or 20µg

of each lipid/well. Additional experimentation was carried out to investigate

fibroblast response to different concentrations of P. gingivalis fDHC. These

experiments were carried out in a similar manner except P. gingivalis fDHC were

studied at 0.2, 2, and 20 µg/35 mm well. After one hour, 30 ng of interleukin-1β (IL-

1β) was added to each sample. After 24 hours, supernatants were harvested and 100

ng of both deuterated prostaglandin E2 (DPGE2) and prostaglandin F2α (DPGF2α)

vi were added. Analyses were performed by derivatization of samples followed by gas chromatograph mass spectrometry (GC-MS). HGF synthesis of PGF2α was unaffected for all lipids tested. IL-1β mediated HGF synthesis of PGE2 was greater for all treatment groups compared to control; however, statistical significance was only achieved for PGDHC and P. gingivalis fDHC (p < .05). P. gingivalis PGDHC and fDHC treated HGF resulted in similar PGE2 synthesis approximately 1.8 fold compared to ethanol vehicle. Morphologic changes suggestive of HGF atrophy and were evident only for PGDHC, and this was noted for the majority of

PGDHC experiments. These findings suggest that IL-1β mediated HGF secretion of

PGE2 can occur even in the absence of a phosphate linkage and 15-carbon isobranched methyl ester, but these structural elements seem to play an important role in the induction of cellular morphological changes observed in phase contrast microscopy.

vii INTRODUCTION

Periodontal diseases include both and periodontitis, which affect the periodontal supporting tissues of teeth [1]. However, unlike gingivitis, periodontitis is accompanied by apical migration of , destruction of connective tissue attachment, and alveolar bone loss [2, 3]. Approximately 30% of the adult U.S. population has moderate periodontitis, with 5% of the adult population experiencing severe periodontitis [4]. In addition, longer average lifespan has resulted in a 32-fold increase in the number of older dentate individuals at risk of developing periodontitis.

More recently, epidemiological studies have shown that may be a significant etiology of various systemic diseases including coronary heart disease, diabetes, and COPD [5].

Periodontal diseases are typically infectious by nature and are a consequence of formation[6-8]. Supragingival biofilm is predominantly composed of Gram- positive aerobic microorganisms and creates and ecological niche enabling Gram negative anaerobic flora to thrive [9]. Specific cultured from periodontal pockets that have been described as more pathogenic than others include Aggregatibacter actinomycetemcomitans, intermedia, , Porphyromonas gingivalis, and denticola where the latter three have been categorized in the red complex of bacteria by Socransky and coworkers [10, 11]. Persistence of these microbes after periodontal therapy has been associated with negative outcomes [12].

Previous investigation has shown that P. gingivalis, a Gram-negative, asaccharolytic, obligate anaerobic rod, is strongly associated with

1 [13]. P. gingivalis produces a broad array of potential virulence factors involved in tissue colonization and destruction as well as in host defense perturbation [14]. Among these

virulence factors, trypsin-like proteinases, called gingipains, have been purified and

characterized, and their functions and pathological roles in periodontitis have been

extensively investigated over the past decade [15]. Gingipains are responsible for most of

the extracellular and cell-bound proteolytic activities produced by P. gingivalis. P.

gingivalis is in close contact with the epithelium in periodontal pockets in vivo [16] and

can invade various cell lines, including epithelial cells [17], endothelial cells [18], and

fibroblasts in vitro [19].

The gingival epithelium is a stratified squamous epithelium that is an interface between the external environment, which is exposed to bacterial challenges, and the underlying periodontal tissue. The basal layer of the gingival epithelium is separated from and attached to the connective tissue by the basal lamina. The gingival epithelium

can be divided into oral, sulcular, and junctional epithelia, based on their architecture.

The , which extends from the oral epithelium to the

facing the teeth, and the junctional epithelium, which mediates the attachment of teeth to

gingiva, are not keratinized, in contrast to the oral gingival epithelium [20]. The sulcular

and the coronal margins of the junctional epithelium are in close contact with bacteria in

the gingival sulcus and appear to be crucial sites with regard to the development of

periodontal diseases. During periodontitis, loss of connective tissue attachment and bone

resorption associated with the formation of periodontal pockets is related to the

pathologic conversion of the junctional and the sulcular epithelium to a pocket

epithelium. Invasion of mammalian epithelial cells is an important strategy developed by

2 to evade the host immune system and cause tissue damage. Gingival

epithelial cells are the primary physical barrier to infections by periodontal pathogens in

vivo. While the epithelium was previously thought to be passive, Dale (2002) [21]

proposed a new perspective, assigning an active role to the epithelium in the host

response to bacterial infections. The epithelium reacts to bacterial challenges by signaling

host responses and integrating innate and acquired immune responses. This review

focuses on the current understanding of host epithelial cells and P. gingivalis interactions

in the pathogenesis of periodontitis. By contrast, periodontally healthy patients will have either no detectable levels of P. gingivalis or only small quantities in subgingival plaque samples [20]. High levels of P. gingivalis have been well correlated with severe

loss of connective tissue attachment, deep periodontal pockets, loss of alveolar bone, and

rapidly progressing periodontal disease [13, 22, 23]. The presence of

periodontopathogenic microorganisms is associated with an increase in inflammatory cell

infiltrate and cytokine production including interleukin-1β (IL-1β), interleukin-6 (IL-6),

interleukin-8 (IL-8), tumor necrosis factor-α (TNF-α), prostaglandin E2 (PGE2), as well

matrix metalloproteinases (MMPs), which recruit lymphocytes, fibroblasts, and

osteoclasts that damage the surrounding periodontal tissues [24].

Subgingival microflora, in general, elicit a complex host immune response

through virulence factors such as endotoxin, proteases, and (LPS).

LPS is a constituent of Gram-negative microorganisms. The LPS molecule

contains three structural components: O-specific polysaccharide, core polysaccharide,

and lipid A. The lipid A portion of the molecule has been shown to be responsible for the

associated biological activity including inflammatory cell activation and inflammatory

3 cytokine production, and its potency differs among different bacterial species [25].

Covalently linked to lipid A by either amide or ester linkages are even carbon-chain 2- hydroxy and odd-chain isobranched 3-hydroxy fatty acids. Bacterial LPS acts by associating with LPS-binding (LBP), and the LPS-LBP complex then binds to

Toll-like receptor-4 (TLR-4) resulting in downstream intracellular signal transduction with activation of nuclear factor-kappa B (NF-κB) and eventual production of proinflammatory cytokines[26, 27]. Toll-like receptors are among a number of pattern recognition surface receptors that appear to play a role in pathogen recognition and initiation of inflammatory and immune responses. Several TLR isoforms exists, and each one has high specificity for binding a certain pathogen-associated molecular pattern

(PAMP) [28]. Evidence has demonstrated that “atypical” P. gingivalis LPS, , and binds TLR-2 receptors [29, 30], and TLR4 appears to be specific for LPS [30].

Bacterial complex lipids recovered in diseased gingival tissues include a class of sphingolipids, called dihydroceramides, and are produced by P. gingivalis [31]. A ceramide is a lipid molecule composed of a sphingosine and a fatty acid. Ceramides are found in high concentrations within the mammalian phospholipid bilayer.

Ceramides have also been key players in cellular response to stress and apoptosis [32].

Hydrolysis of sphingomyelin by acid sphingomyelinase results in the release of ceramide,

while synthesis of ceramide is catalyzed by the serine-palmitoyl-CoA transferase and ceramide-synthase [33]. Sphingomyelin is a type of sphingolipid found in mammalian cell membranes most significantly in the myelin sheath around certain nerve cell axons.

It usually consists of phosphorylcholine and ceramide. Sphingomyelin is an important

4 facilitator of signal transduction. Sphingolipids are derived from the aliphatic amino alcohol sphingosine and play an important role in both signal transmission and cell recognition. Ceramide molecules reorganize the cell membrane leading to the formation of large, distinct ceramide-enriched membrane domains, termed rafts, that serve to cluster and aggregate activated receptor molecules [34]. Besides the important role of ceramide in receptor-induced cell death, several studies demonstrate a central role of ceramide in

UV-light, irradiation and triggered cell death [35]. Several recent publications indicate that ceramide is also present in mitochondria, and that mitochondrial ceramide may play a role in apoptosis. Mitochondrial ceramide seems to be generated through the activity of the acid sphingomyelinase, which has been shown to reside in the space between the inner and outer mitochondrial membrane [36]. Although many studies indicate a central role of ceramide for the induction of apoptosis after an acute stimulus, only a few data exist that demonstrate a contribution of ceramide to a dysregulation of apoptosis in human disorders. Diseases that have been shown to be caused by dysregulation of cellular ceramide include neurological syndromes, including hereditary sensoric neuropathy Type

I and II. Hereditary sensoric neuropathy Type I is caused by a mutation of serine– palmitoyltransferase 1 and an increased cellular ceramide concentration, which seems to be causally linked to degeneration of peripheral sensory neurons [37, 38]. Cellular ceramide levels are also increased in the Batten’s syndrome, which is caused by a mutation of palmitoyl-thioesterase and of CLN3 resulting in neuronal cell death with cognitive and motor decline, seizures and blindness [39]. Finally, experiments on

Drosophila also suggest an involvement of ceramide in retina degeneration, although it is

5 not known whether similar mechanisms are effective in the corresponding human disease

[40, 41].

Ceramides have been linked to periodontal pathogenesis since they are readily recovered from disease gingival tissues and root surfaces of hopeless teeth extracted due to severe periodontitis [42]. More specifically, P. gingivalis, dihydroceramides have been detected in lipid extracts of -contaminated teeth, subgingival calculus, subgingival plaque and in diseased gingival tissues. This evidence suggests that diseased gingival tissues are not simply contaminated with subgingival plaque. Regardless of the processes resulting in bacterial lipid accumulation in periodontitis tissue samples or metabolic clearance of these bacterial lipids, the recovery of phosphorylated dihydroceramides in diseased gingival tissues indicates that the biological properties of the contaminating bacterial lipids must be understood within the context of disease promotion [42].

Sharing structural similarity to lipid A, previous chemical analysis via gas chromatography-mass spectrometry (GC-MS) has shown that these bacterial ceramide lipids also contains 3-hydroxy fatty acids linked to a long chain base of either 2-amino,3- octadecanediol or 2-amino1,3-nondecanediol amide linkage [31]. Bacterial ceramides share basic structural characteristics with mammalian ceramides and are therefore expected to promote cell activation in host cells normally susceptible to elevated levels of mammalian ceramides [43, 44]. P. gingivalis synthesizes free, phosphoethanolamine, and phosphoglycerol dihydroceramides that share a core lipid structure consisting of three long chain bases (17, 18, and 19 carbon aliphatic chains) in amide linkage to 3-OH isobranched C17:O [45, 46]. The phosphoglycerol dihydroceramide class may be

6 substituted with isobranched C15:O linked to the beta hydroxyl of 3-OH isobranched C17:O

[46]. These bacterial lipids are not components of lipopolysaccharide and their recovery

in gingival tissues is not attributable to coincident bacterial invasion [31, 45]. In

addition, mammalian cells have not been shown to produce 3-OH isobranched C17:O, and

this fatty acid has been recovered in lipid extracts of periodontally diseased human

and gingival tissues [45].

P. gingivalis dihydroceramides can be extracted with organic solvents directly from the bacterial pellet obtained after growth of the organism, centrifugation, and lyophillization following a protocol described by Bligh and Dyer, and Garbus et al [47,

48]. Following total lipid extraction, select dihydroceramides are separated by direct and reverse phase high performance liquid chromatography (HPLC) [46]. Confirmation of

these fractions is possible with electrospray (ESI-MS) or gas chromatograph mass spectrometry (GC-MS) prior to experimentation [46].

For many years, periodontal therapy only aimed to remove the etiologic factors

from the root surface and recontour hard tissue anatomical defects. This approach

minimizes periodontal and creates a clinical situation that is more easily

accessible for home and professional practices. With advances in tissue

engineering and the importance of esthetics, regeneration of the periodontal attachment

apparatus has been considered the ultimate goal of periodontal treatment. The

is a complex organ comprised of gingiva, periodontal ligament, alveolar

bone, and cementum. To achieve successful periodontal regeneration, formation of an

epithelial seal, insertion of new connective tissue fibers into the root, reformation of a

new acellular cementum reformed on the root surface, and restoration of alveolar bone

7 are required [49]. However, the inhibition of periodontal regeneration after conventional

treatment modalities is due to proliferation of epithelial tissues into the periodontal defect

at a faster rate than mesenchymal tissues [50]. This leads to the formation of a long

junctional epithelium and prevents the selective migration, proliferation, and

differentiation of cells derived from the periodontal ligament (PDL) [51]. The concept

that only PDL cells have the potential to create a new connective tissue attachment has

been confirmed through investigation giving the basis for the clinical application of

guided tissue regeneration (GTR) [52]. However, the results obtained using the GTR

technique are reported to be unpredictable and of small magnitude [53, 54] possibly because of postoperative infections and the inability to completely seal the healing area

[55]. Three basic components must be considered in periodontal tissue engineering: 1) appropriate signals, 2) scaffolds/barriers to enable coronal population of the appropriate cells from the PDL, and 3) cells from the PDL with regenerative potential. It has been shown that PDL cells may prevent ankylosis and root resorption in addition to regeneration of a new periodontal ligament with insertion into both tooth and bone. Also, roots covered with osteoprogenitor cells induce cementum-like tissue formation suggesting that cementoblast and osteoblast precursors commonly originate from the alveolar bone [56, 57]. The formation of new bone and cementum varies from complete absence to a thin layer of newly formed cementum with complete new bone formation covering the entire previously denuded root surface. Moreover, it may be speculated that differentiation of PDL cells is highly sensitive to differences in their microenvironments, resulting in different types of periodontal healing. It has been shown that, [58] after transplantation, the number of cells increases over time and reflects sustained

8 proliferation. These newly transplanted cells migrate to the bone surface and express

osteoblast markers, for example osteopontin and bone sialoprotein, to facilitate

osteogenesis; cells that migrate to PDL express the same phenotypic markers as other

PDL cells. This confirms the hypothesis that, under some circumstances, PDL cells can

promote periodontal regeneration.

Independent of the periodontal disease status, the predominant cell type recovered

in gingival connective tissue is the fibroblast [59]. Fibroblasts play an important role in

the remodeling of extracellular matrix (ECM) by synthesizing connective tissue

components predominantly constituted of fibrillar collagens [60]. Fibroblasts respond to

various inflammatory cytokines and growth factors [61], and control the balance between

ECM synthesis and degradation [62]. A shift in this normal homeostasis can lead to direct destruction of the periodontal connective tissue attachment apparatus and indirect destruction of alveolar bone through increased synthesis and secretion of proinflammatory cytokines (PGE2, IL-1β, IL-6, IL-8, and TNF-α) [63]. Gingival fibroblasts exposure to bacterial lipids is expected given the impressive recovery of bacterial lipids in diseased periodontal tissues [45, 64]. Fibroblasts are capable of secreting high levels of prostaglandin E2 (PGE2) [65], a proinflammatory host factor with ability to have adverse effects on bone and connective tissue homeostasis.

Previous investigation has described methods regarding human gingival fibroblast culture from gingival tissues harvested during periodontal surgery [65]. Once cultures have been obtained, one can apply fibroblasts to wells contaminated with dihydroceramides, introduce IL-1β, and measure response via prostanoid quantification

9 [46]. Methods of separation, derivatization, and chemical analysis of prostanoids have

also been previously described [66-68].

Prostaglandins are a subclass of eicosanoids, derived from arachidonic acid, and are important mediators of inflammation. Prostaglandin production, more specifically

PGE2, has been strongly associated with periodontal pathogenesis [69]. Most notably, of

their many functions, prostaglandins are potent vasodilators and inducers of cytokine

production by various cells. In conjunction with proinflammatory cytokines, PGE2 acts on fibroblasts and osteoclasts, to induce MMP production, which is also a major component to the destructive process of periodontitis [69]. As demonstrated in a murine model, PGE2 also appears to significantly facilitate osteoclastogenesis by enhancing

RANKL (receptor activator of nuclear factor-kappa B ligand) and depression of

osteoprotegerin (RANKL inhibitor) [70]. Therefore, PGE2 association with destruction

of connective tissue attachment and alveolar bone supports its central importance in the

periodontal disease process. Previous investigation has associated PGE2 with

periodontitis and suggests that its concentration in gingival crevicular fluid increases in

gingivitis relative to health and is at very high concentration during periods of

periodontal disease progression [71].

Previous investigation has shown prostaglandin hypersecretory response of

gingival fibroblasts to both phosphorylated dihydroceramide classes in vitro [46].

Phosphoglycerol dihydroceramides cause the most significant morphologic changes in

gingival fibroblasts suggesting either apoptosis and/or cell necrosis. This line of

evidence demonstrates the biological activity of these lipids and supports their inclusion

among several microbial virulence factors of P. gingivalis [46]. Interestingly, short chain

10 ceramides have been shown to promote interleukin-1-mediated prostaglandin E2 secretion from dermal fibroblasts [72], and COX-1 in fibroblasts [73]. However, specific structure-function relationships of P. gingivalis dihydroceramides have yet to be determined.

11

OBJECTIVE

The aim of this investigation was to isolate P. gingivalis phosphoglycerol dihydroceramides (PGDHC), unsubstituted PGDHC (PGDHC-C15:0), free DHC (fDHC), synthetic fDHC (A) and dephosphorylated PGDHC, and correlate structure with biological activity based on human gingival fibroblast (HGF) prostaglandin secretory response in vitro.

12

MATERIALS AND METHODS

Lipid extraction from P. gingivalis

Previously, P. gingivalis (ATCC 33277, type strain) was inoculated into cooked medium supplemented with trypticase, yeast extract, menadione, and hemin (BBL,

#295982, Becton Dickinson, Franklin Lakes, NJ). Once growth was established, the initial culture was inoculated into enriched thioglycollate medium (BBL) and prereduced blood agar plates. Batch suspension cultures were established after verification of anaerobic growth of Gram-negative rods in thioglycollate medium and demonstration of black-pigmented bacterial colonies on blood agar plates under anaerobic culture conditions. Batch suspension cultures were grown in basal [peptone, trypticase, and yeast extract, (BBL)] medium supplemented with hemin and menadione (Sigma; St. Louis,

MO) and brain heart infusion. The suspension cultures were incubated in an anaerobic chamber flushed with N2 (80%), CO2 (10%), and H2 (10%) at 37°C for five days, and the bacteria were harvested by centrifugation (3,000 g for 20 min). Following lyophilization,

~4 g of P. gingivalis pellet was extracted overnight using a modification of the phospholipid extraction procedures of Bligh and Dyer [47] and Garbus et al. [48].

Specifically, 4 ml of H20 + 16 ml of MeOH-CHCl3 (2:1; v/v) was added to the bacterial sample and vortexed. After 12 h, 3 ml of 2 N KCl + 0.5 M K2HPO4 and 3 ml CHCl3 were added and the sample vortexed. The lower organic phase was carefully removed and

CHCl3 (3 ml) was added to each sample and vortexed. The CHCl3 phase was removed

13 and combined with the previous organic solvent extract. The lipid extract from P. gingivalis was dried under nitrogen and stored frozen. This sample of frozen P. gingivalis lipid extract was donated as a kind gift from Dr. Frank Nichols.

Lipid fractionation by HPLC

Direct phase HPLC (d-HPLC) [1 x 25 cm silica gel, 5 µm; Supelco, Inc., Bellefonte, PA] was used for initial separation of bacterial lipids as well as repurification using hexane- isopropanol-water (6:8:0.75; v/v/v: solvent A) with isocratic elution [74]. Reverse phase

HPLC (r-HPLC) [250/4mm diatomaceous earth, 5 µm; Phenomenex] was used for further purification of fDHC using propanol:acetontrile (2:3; v/v: solvent B) with isocratic elution as well. In preparation for d-HPLC, lipid samples were dissolved in solvent A to achieve a concentration of ~80 mg/ml. For each chromatographic separation,

~3 mg of lipid was applied and fractions were pooled for 25 column fractionations.

Samples were eluted at 2.0 ml/min with 1 min fractions collected. For r-HPLC, lipid samples were dissolved in solvent B. For each chromatographic separation, ~1 mg of lipid was applied and fractions 1-25 were collected. During fractionation, lipid samples were continuously monitored at 205 nm with a UV-vis spectrophotometer. Fractions were dried under nitrogen and stored at resuspended in 2 ml of solvent A.

14

Electrospray mass spectral analysis

An additional 10 µl from each fraction was analyzed for dihydroceramide constituents using electrospray-MS (ESI-MS). Electrospray mass spectrometry analysis was accomplished using a Micromass Quattro II mass spectrometer system. Dihydroceramide lipid fractions, dissolved in solvent A, were injected at a maximum concentration of 100

µg/ml. Lipid samples (10 µl) were infused at a flow rate of 20 µl/min. For electrospray positive-ion analyses, the desolvation and inlet block temperatures were 100°C and

150°C, respectively, and the transcapillary potential was 3,500 volts. For electrospray negative-ion analyses, the desolvation and inlet block temperatures were 80°C and

100°C, respectively, and the transcapillary potential was 3,000 volts. The cone voltage was usually 30 volts, and the mass acquisition range was 0–2,000 amu for initial electrospray MS analyses. Fractions containing specific dihydroceramide lipids were pooled and repurified using the same column and elution conditions. The repurified lipids were evaluated for dihydroceramide recovery using ESI-MS. A purified, synthetic fDHC

(A) was obtained from the University of Connecticut Storrs Department of Chemistry to include in this investigation.

Preparation of test samples

Lipid recovery in each HPLC fraction was determined by drying 5 µl from each fraction onto a microbalance tray and weighing the tray using a Cahn Electrobalance (Ventron

15 Corporation, Cerritos, CA). Predetermined amounts of PGDHC, PGDHC-C15:0, A, and

fDHC were transferred in CHCl3 to conical vials and dried. These selected HPLC

fractions were then dissolved in ethanol to achieve a final concentration of 0.25 µg/µl,

and 80 µl of each lipid fraction was deposited in 35 mm culture wells while the culture

dishes were maintained in a laminar flow hood. The ethanol was allowed to evaporate

overnight, leaving a lipid residue covering most of the culture well surface. Vehicle

control wells received only ethanol solvent. Lipids were deposited onto culture dishes in

this manner so that cells could be exposed to bacterial lipid fractions without introducing

organic solvent into the culture medium. In a second series of experiments, fDHC was

diluted to 0.0025 µg/µl, 0.025 µg/µl and 0.25 µg/µl, and 80 µl were deposited into each

respective 35 mm well for dose response comparison. PGDHC, PGDHC-C15:O, and A were also examined in this second series of experiments.

Determination of biological activity with human gingival fibroblasts

Primary cultures of gingival fibroblasts were obtained using gingival tissue, harvested at the time of periodontal surgery, from two separate patients and processed as previously described [65]. Gingival tissue samples were obtained according to a protocol approved by the University of Connecticut Health Center Institutional Review Board, and participants provided written informed consent. Primary cultures of gingival fibroblasts were grown in Minimal Essential Medium (MEM) supplemented with 10% fetal calf , penicillin G (100 U/ml), streptomycin (100 µg/ml), and amphotericin B (0.25

µg/ml) (Sigma), and cells were passaged fewer than 10 times for these experiments. The

16 cells were harvested using trypsin (0.1%, w/v) in Ca2+-, Mg2+-free phosphate-buffered saline and resuspended in MEM to achieve a 1:3 dilution of cells for passaging. Cells were cultured in an H2O-saturated, 5% CO2 atmosphere at 37°C. Fibroblasts were

introduced to bacterial lipids by inoculation into culture wells to achieve a cell

number/surface area equivalent to confluent cultures. After 1 h of exposure to bacterial

lipids, 30 µl of culture medium containing human recombinant IL-1ß (final concentration,10 ng/ml) (Immunex; Seattle, WA) was injected to each sample. After the cultures were incubated for an additional 24 h, they were photographed at x100 magnification using an Olympus phase contrast light microscope. Photomicrographs of the effects of P. gingivalis total lipid sample were included from a pilot experiment.

However, quantification of total lipid induced prostaglandin production was not included in this experiment. Medium samples were then harvested and frozen.

Quantification of prostaglandin production by GC-MS

Prostaglandin E2 (PGE2), and prostaglandin F2α (PGF2α) were quantified in medium

samples using a modification of the method of Luderer et al. [66], previously described

by Nichols [67]. Each medium sample was supplemented with 100 ng of D4- PGF2α and

D4-PGE2. The fibroblast medium samples were thawed and the pH adjusted to pH 3.5

with concentrated . The acidified medium samples were then applied to

reverse-phase preparative columns [Supelclean, LC-18 SPE tubes, 6 ml; Supelco, Inc.] mounted on a vacuum manifold. Before applying the medium samples, the C-18 columns

(3 ml) were regenerated by running sequentially 3 ml of 100% methanol followed by 3

17 ml of Ca2+-Mg2+-free phosphate-buffered saline (pH 3.5). Each acidified medium sample was then applied to individual SPE columns, the columns were washed with 3 ml of 25% methanol in water, and enriched prostaglandins were eluted with 3 ml of 100% methanol. The resultant samples were then supplemented with 2 ml of 1% formic acid in water, and the prostaglandins were extracted with 2 ml of chloroform. The chloroform extracts were dried under nitrogen.

All derivatizing agents were obtained from Pierce Chemical Corp. (Rockford, IL).

Prostaglandin samples were derivatized using the method of Waddell, Blair, and Wellby

[68]. Prostaglandin samples were first treated with 2% methoxylamine hydrochloride in pyridine (30 µl). After standing overnight at room temperature, the samples were dried under nitrogen, dissolved in acetonitrile (30 µl), and treated with pentafluorobenzyl bromide (35% v/v in acetonitrile, 10 µl) and diisopropylethylamine (10 µl). The samples were vortexed, incubated for 20 min at 40°C, and evaporated under nitrogen. The residue was then treated with bistrimethylsilyl-trifluoroacetamide (BSTFA, 50 µl) and allowed to stand at room temperature for 4–5 days.

Derivatized products of hydroxy fatty acids, long-chain bases, and prostaglandins were transferred to vials used for GC-MS automated sampling (crimp cap vials, 200 µl). GC-

MS was carried out on a Hewlett Packard (Avondale, PA) 5890-gas chromatograph interfaced with a 5988A-mass spectrometer. Prostaglandin samples were applied to an

SPB-1 (12 m x 0.2 mm, 0.33 µm film thickness; Supelco, Inc.) column held at 100°C.

Prostanoid samples were analyzed using a temperature program of 5°C/min from 100°C to 240°C. The injector block was held at 260°C, and the transfer tube was maintained at

280°C. Prostaglandin levels were quantified using selected ion monitoring of the

18 characteristic base peak ions, and bacterial fatty acids and long-chain bases were detected and interpreted through analysis of characteristic spectra.

NMR analysis of fDHC

NMR data were collected on a Bruker (Rheinstetten, Germany) DRX-400, a Bruker

Avance 500, and a Varian (Palo Alto, CA) INOVA 600 (operating frequencies 400.144

MHz, 500.13 MHz, and 599.75 MHz, respectively). For bacterial fDHC, HPLC fractions

14 and 15 lipids dissolved in a mixture of deuterated solvents (CDCl3-CD3OD). These

data allowed the assignment of proton resonances of the dihydroceramide base and the

phosphate side-chain of the molecule, as well as specific components of the aliphatic

chains. A mixture of solvents was necessary in order to prevent aggregation into micelles

or reverse micelles, depending on the solvent. Synthetic fDHC (A) was aliquoted,

dissolved in a mixture of deuterated solvents (CDCl3-CD3OD) and analyzed in a similar

fashion. This further allowed for comparison between synthetic (A) and P. gingivalis

fDHC.

Statistical Analysis

Test groups were compared to ethanol vehicle and each other using analysis of

variance (ANOVA) after calculation of test/control (T/C) ratios. Fisher PLSD, Scheffe

F-test and Dunnett t were used to determine significance of ANOVA at 95% confidence

19 level. Statistical analysis was computed using a computer software package for

Macintosh (StatView).

20 RESULTS

P. gingivalis bacterial complex lipid structures with corresponding mass to charge

ratio (m/z) including PGDHC, PGDHC-C15:0, and fDHC can be seen in (Fig. 1). Lipids were extracted from P. gingivalis and separated by dHPLC and rHPLC. Several dihydroceramide lipid moieties can be observed in the ESI-MS spectrum for the P. gingivalis total lipid sample in (Fig. 2). Samples of PGDHC were obtained by dHPLC separation of the total lipid extract, and purity of samples was confirmed with ESI-MS analysis (Fig. 3). PGDHC-C15:0 was obtained by NaOCH3 treatment of an aliquoted

sample of PGDHC, separated by dHPLC, and purity of the samples was confirmed with

ESI-MS analysis (Fig. 4). High purity of samples can be noted by comparison of relative

ion abundance of m/z peaks corresponding to each separated dihydroceramide.

Corresponding solvent ESI-/+M spectrum is located below the separated

dihydroceramide spectrum in each figure.

Pure samples of fDHC were obtained using rHPLC. A sample of a UV-vis

spectrum for fDHC fractionation after initial rHPLC separation is shown in Fig. 5 where

fDHC elution from column is delayed and can be observed in the later peak. After

collection and pooling of these corresponding fractions, rHPLC was used a second time

(Fig. 6). In this UV-vis spectrum, the largest signal corresponds with refined separation

of fDHC. These fractions containing fDHC were analyzed with ESI+MS and this mass

spectrum can be seen in Fig. 7. As previously described, synthetic fDHC was provided

by the University of Connecticut Storrs Department of Chemistry. After synthesis and

purification, characterization was performed similarly with addition of NMR

21 spectroscopic analysis. ESI+MS for synthetic dihydroceramide (A) can be seen in Fig. 8.

NMR spectra for both fDHC and A can be observed together for comparison in Fig. 9.

An impressive similarity is noted including the finger print region between both spectra.

The ESI+MS and the NMR data provide strong evidence of structural similarity between

fDHC and A.

Morphologic changes of human gingival fibroblasts (HGF) were evaluated after

24 hours of exposure to lipid or ethanol vehicle in addition to 30 ng IL-1ß in 35mm wells with phase contrast microscopy. Photomicrographs for HGF exposure can be observed in

Fig. 10. Morphologic changes could not be observed for ethanol vehicle (Panel A) and/or fDHC (Panel D). However, morphologic changes appearing as severe atrophy and apoptosis were observed after 24 hours of exposure to 20µg of PGDHC + 30 ng IL-

1ß (Panel C) and 20µg Total Lipid extract + 30 ng IL-1ß (Panel B) in 35mm wells.

Treatment of HGF with P. gingivalis PGDHC, PGDHC-C15:0, and fDHC as well as synthetic free DHC demonstrated varying potentiation of IL-1ß mediated PGE2

secretion for both 20µg dose and fDHC dose response of 0.2, 2.0, and 20µg when

compared to vehicle control. PGF2α secretion was unaffected by bacterial and synthetic

lipids (Fig. 11). In a separate experiment comparing the effects of PGDHC-C15:0, a trend

suggesting stimulation of HGF PGE2 secretion was noted for all lipids, but statistical

significance was not achieved (Fig. 12). Although trends were noted for stimulation for

all treatment groups compared to control, statistical significance was only noted for

PGDHC (20µg ) and fDHC (20µg ) (Fig 13). A trend for increased PGE2 secretion with

increasing concentration of fDHC was also noted, but did not achieve statistical

22 significance (Fig. 14). The effects of all samples on HGF PGE2 and PGF2α secretion was determined by calculating treatment/control (T/C) for Figures 12-14.

23 DISCUSSION

This present investigation demonstrates that, in vitro, P. gingivalis PGDHC,

PGDHC-C15:0, fDHC, and synthetic fDHC differentially induce IL-1β mediated HGF

secretion of PGE2. HGF secretion of PGF2α was unaffected by P. gingivalis and

synthetic dihydroceramides. PGDHC and fDHC treated HGF resulted in similar PGE2

synthesis approximately 1.8 fold compared to ethanol vehicle (p < .05).

Several proinflammatory cytokines including IL-1β [75], tumor necrosis factor α

[76], epidermal growth factor [77], and transforming growth factor β [78] are known to

stimulate PGE2 production. Prostaglandin synthesis is regulated by the activity of both

phospholipase A2 and cyclooxygenase (COX) [79]. Previous investigation has elucidated

that the rate-limiting step to prostaglandin synthesis is related to COX stability since it is

rapidly degraded and has a short half-life [80]. Adding to the complexity of this

mechanism, IL-1 induced COX synthesis is mediated by protein-dependent post-

transcriptional events [81, 82]. Progressively, in vitro, examination of human dermal

fibroblast IL-1-mediated PGE2 secretion has determined that spingosine and C2- ceramide treatment resulted in an 8-fold induction of COX mRNA within 1-2 hours.

Although lacking a phosphate head group and isobranched fatty acid linkage, C2 ceramide has a structure relatively similar to P. gingivalis dihydroceramides. Given these findings, as found for human dermal fibroblasts, a similar mechanism is expected where

P. gingivalis and synthetic lipid samples tested in this study may increase COX expression accounting for the observed enhancement of IL-1β mediated PGE2

production.

24 More specifically, increased sulcular concentration of IL-1β and PGE2 has been

previously shown to regulate the magnitude of tissue destruction in progressive

periodontitis [83-88]. IL-1 is a proinflammatory cytokine that has a large array of

biological activities and directly regulates several genes expressed during inflammation

[89]. There are two principal forms of IL-1 with agonist activity: IL-1α and IL-1β. A third ligand, IL-1ra, is an antagonist that functions as a competitive inhibitor. Two IL-1 receptors are found on the surface of target cells: IL-1 receptor-1 and IL-1 receptor-2.

IL-1 is produced by many cell types including monocytes/macrophages, neutrophils, fibroblasts (gingival and periodontal ligament), epithelial cells, endothelial cells, and osteoblasts [90-93], and can induce upregulation of adhesion molecules on leukocytes and endothelial cells, stimulate production of chemokines, increase production of prostaglandins, matrix metalloproteinases, and even enhance bacterial killing and phagocytic activity [89, 94]. There are several lines of evidence that implicate IL-1 in the pathogenesis of periodontal disease [95-97]. Furthermore, experimental evidence suggests that susceptibility to periodontal disease is influenced by genetic polymorphisms of the IL-1 gene [98, 99]. Interestingly, IL-1 gene polymorphisms have also been associated with poorer periodontal prognosis in smokers where risk of tooth loss increased up to 7.7 times over a 14-year periodontal maintenance period [100].

Impressive levels of IL-1 have been recovered in gingival crevicular fluid (GCF) in patients with periodontal disease compared to healthy controls. According to Salvi et al.,

GCF IL-1 levels were approximately 17 ng/ml for healthy controls, 116 ng/ml for gingivitis patients, 263 ng/ml for chronic periodontitis patients and as high as 458 ng/ml for patients [101]. Similarly, many studies have reported that

25 GCF PGE2 levels are significantly elevated in patients with periodontal disease compared

to healthy controls. In a monkey model, previous investigation has shown that crevicular

fluid concentration of PGE2 may be elevated up to threefold during only two months of

periodontitis progression [83]. In the same model, with a placebo-controlled double-

blinded study, this group has also demonstrated that local administration of a non-

steroidal anti-inflammatory agent (Ketoprofen cream) significantly reduced crevicular

fluid PGE2 concentration with corresponding gain in alveolar bone over a 6 months follow up period. Confirmed in a human model of experimental gingivitis, increased

GCF PGE2 levels were found at 4 weeks following cessation of oral hygiene procedures

[85]. According to Seymour et. al., GCF PGE2 levels (ng/ml) were as high 20, 50, 42,

82, and 90 examining 21 healthy, 28 gingivitis, 21 periodontitis, 15 aggressive

periodontitis, and 15 refractory periodontitis patients, respectively [84]. In review of

these findings, it seems clear that IL-1 and PGE2 are highly important inflammatory mediators implicated in the pathogenesis of periodontitis.

Previous literature has demonstrated that 3-OH isoC17:0 is readily recovered in

both organic and aqueous solvent extracts of plaque samples, teeth and root sections

substantiating the presence of bacterial complex lipid and LPS levels, respectively. This

bacterial hydroxy fatty acid is only recovered from lipid extracts of P. gingivalis,

Prevotella, Bacteroides, [102, 103], Flavobacterium [104, 105], and

Myxobacteria [106], but not synthesized by mammalian tissues.

These findings are similar to results found previously examining the effects of P.

gingivalis total lipid extract and PGDHC on HGF cultures, in vitro [46]. However, in this

study, less impressive recovery of PGE2 was noted. This finding may be a result of

26 response variation between subjects or associated with prolonged age of the cell cultures.

Genetic variation of fibroblast phenotype, although not related to secretion of

prostanoids, has been exhaustively demonstrated in association with hereditary gingival

fibromatosis [107]. Given this information, one could conclude that it is quite possible to

have a broad range of HGF responsiveness when exposed to P. gingivalis lipids.

Although HGF were passaged less than ten times, this treatment occurred over a period

of approximately eight months. A more representative HGF synthesis of PGE2 would be expected with increased sample size of subjects for biopsy and decreased storage time of

HGF clones. In consideration of these limitations, future investigations will include increased numbers of subjects willing to donate biopsy samples as well as completion of testing within a shorter time frame.

LPS is a devastating of P. gingivalis, Aggregatibacter

actinomycetemcomitans, as well as other Gram-negative bacteria. LPS acts as a microbe-

associated molecular pattern recognized through pattern-recognition receptors on resident

immune and non-immune cells within the periodontium [108]. As a result, inflammatory

cells are recruited, osteoclasts are activated, and prostanoids, lytic enzymes, and

proinflammatory cytokines are secreted [109, 110]. Murine models have been used to demonstrate the devastating effects localized LPS injection has on the periodontium including severe inflammation and alveolar bone loss. Historically, these studies isolated

LPS from non-periopathogenic bacteria including Escherichia coli or Salmonella typhimurium [111-113]. However, more recently, Rogers et al. observed that local injection of A. actinomycetemcomitans LPS in Sprague-Dawley rats induced severe bone

27 loss over an 8 week period using micro-computed tomography and histologic evaluation

[114].

The available literature clarifies the virulence potential of both bacterial complex

lipids and LPS to play a direct role in periodontal pathogenesis. Although both of these

lipid groups are components of the bilayer membrane of certain bacteria involved in

periodontal disease, controversial evidence suggests that the recovery of LPS in diseased

gingival tissues may be less than expected. As stated previously, long-chain 3-hydroxy fatty acids are covalently linked to the lipid A component of LPS as well as the long chain base of bacterial complex lipids of P. gingivalis, Prevotella, Bacteroides,

Capnocytophaga [102, 103], Flavobacterium [104, 105], and Myxobacteria [106], but

not synthesized by mammalian tissues. After extraction procedures, Nichols (1994)

found that LPS 3-hydroxy iC17:0 and bacterial complex lipid 3-hydroxy iC17:0 are

recovered in the aqueous and organic phase, respectively [45]. Subsequently, both LPS

and bacterial complex lipid distribution were determined for two periodontal pathogens

(P. gingivalis and ), subgingival plaque samples and gingival tissue

samples both isolated from gingivitis and severe periodontitis sites. Mean values of 3-

hydroxy iC17:0 recovered in organic solvent extracts of P. gingivalis and P. intermedia

strains ranged from 56-63% and only < 5% of the total 3-hydroxy iC17:0, respectively.

Organic solvent extract of subgingival plaque samples contained 75% of the total 3-

hydroxy iC17:0 for periodontitis subjects and only 43% for gingivitis subjects. Finally, for

gingival tissue samples, 3-hydroxy iC17:0 was primarily recovered only in organic solvent extracts of both healthy and mildly inflamed and periodontitis gingival tissue samples. In conclusion, these results indicate that LPS from these organisms is not present in

28 substantial levels in diseased gingival tissues. Furthermore, the significant recovery of 3- hydroxy iC17:0 in organic solvent from periodontitis plaque samples and gingival tissue suggests that bacterial complex lipid, from the aforementioned select bacterial species, may be responsible for a more significant fraction of the lipid component of periodontal pathogenesis.

Of clinical relevance, adequate removal of lipids for experimental purposes requires the use of toxic organic solvents such as chloroform [42]. Since these structures are extremely hydrophobic, their removal during routine non-surgical and surgical therapy will be limited to mechanical of cementum and locally inflamed soft tissue. Significant amounts of lipid remaining on the root surface are expected since they are insoluble by aqueous irrigation, and mechanical debridement of teeth to remove calculus is limited even with adequate surgical access [115]. Removal of bacterial complex lipids in inflamed gingival tissues with resective periodontal surgery may be related to the observation of decreased units of gingival inflammation, evident in 6 month follow-up biopsy samples, compared to non-resective [116]. Furthermore, the persistence of lipid on the root surface may also have limiting effects on regeneration potential of the periodontium. Not only were the HGF significantly reactive to the presence of both PGDHC and P. gingivalis fDHC, as evident by secretion of PGE2, but the majority of photomicrographs taken of HGF exposed to PGDHC revealed cellular morphological changes suggestive of apoptosis.

Of the bacterial lipids and synthetic fDHC examined, HGF morphologic changes were only noted for PGDHC; additionally, the majority of PGDHC experiments resulted in appearance of cellular changes observed with phase contrast microscopy. A potential

29 explanation for this finding may be the presence of both a phosphate head group and an

ester-linked 3-OH 15-carbon isobranched fatty acid. This unique combination of these

structural elements of PGDHC may be specific for the molecule’s biologic activity on

HGF in vitro. Interestingly, LPS lipid A contains two phosphate groups coupled to

glucosamines, and previous investigation has shown that removal of one of these

phosphate groups generates a monophosphoryl lipid A that is 100-fold less toxic than the

unmodified lipid A [117]. Alkaline phosphatases have been shown to modify LPS by

dephosphorylating its lipid A moiety [118-121]. is a glycoprotein

and membrane bound enzyme that increases local concentrations of phosphate ions [122].

In the periodontium, alkaline phosphatase is a very important enzyme as it is part of the

normal turnover of the periodontal ligament, root cementum formation and maintenance,

and bone homeostasis [122]. It is produced by fibroblasts, osteoblasts, osteoclasts, as

well as many other cells, but the primary source of alkaline phosphatase in gingival

crevicular fluid is neutrophils [122] where it functions in superoxide generation, an

important defense mechanism of this cell [123]. In a more recent investigation, using a

zebrafish model, Guillemin and coworkers have demonstrated the LPS detoxifying

effects of alkaline phosphatase [124]. No effect on survival was noted when wild-type

animals were exposed to LPS and when animals with blocked expression of intestinal

alkaline phosphatase were exposed to previously dephosphorylated LPS via calf intestinal alkaline phosphatase. However, when animals with blocked expression of intestinal alkaline phosphatase were exposed to LPS, toxic effects were readily apparent with 100% mortality within the following 48-hour time period after administration. These authors conclude that intestinal alkaline phosphatase protects these animals from the toxic effects

30 of LPS, and that “commensal” flora may actually be toxic in the absence of this adaptive

physiology. In this experiment, bacterial fDHC and PGDHC stimulated IL-1β HGF

PGE2 secretion was similar despite the absence of the phosphate head group in the fDHC

organic structure. Aside from the minimal recovery of LPS from plaque and gingival

tissue samples from patients with periodontitis as mentioned previously, possible

protection of LPS toxicity by GCF alkaline phosphatase dephosphorylation warrants

further investigation. Future experimentation examining the effects of bacterial complex

lipid and LPS phosphate groups may lead to a better understanding of structure function

relationships and the role of alkaline phosphatase in periodontal disease mechanisms.

31

CONCLUSION

Further studies are needed to evaluate the structure function relationship of

bacterial complex lipids and LPS as well as the potential protective effect of alkaline

phosphatase in periodontal diseases. Larger sample sizes may clarify the potential effects

of these various structural elements as well as dose response relationships. Other future

investigations including effects on osseointegration, development of multiple sclerosis in

a murine model, and the removal of lipids from contaminated root surfaces are either

planned or currently in progress.

In summary, through the use of reverse and direct phase HPLC, pure samples of

P. gingivalis PGDHC, PGDHC-C15:0, and fDHC were obtained and confirmed with ESI-

MS (ESI+MS for fDHC). Synthetic fDHC was obtained from the University of

Connecticut Storrs graduate chemistry department, and molecular structure was confirmed using ESI+MS and NMR with consistent comparison to P. gingivalis fDHC.

In vitro, P. gingivalis PGDHC, PGDHC-C15:0, fDHC, and synthetic fDHC differentially

induce IL1-β mediated HGF secretion of PGE2. HGF secretion of PGF2α was unaffected

by P. gingivalis and synthetic dihydroceramides. PGDHC and fDHC treated HGF

resulted in similarly increased PGE2 synthesis approximately 1.8 fold compared to

ethanol vehicle (p < .05). In addition, morphologic changes suggestive of HGF atrophy

and apoptosis were evident only for PGDHC, and this was noted for the majority of

PGDHC experiments.

32

Phosphoethanolamine Iso C15:0-substituted Unsubstitu ted Phosphatidyl- Dihydroceramides Phosphoglycerol Phosphoglycerol Free Dihydroceramides ethanolamines (PE DHC) Dihydroceramides Dihydroceramides (PEA) (PG DHC) (Unsub PG DHC) (Free DHC) H H OH OH H H H N H N OH OH Phosphorylated Head HO HO Groups O O O O O O- O O- O P O P P O P O O O O HO O H H H H N N N N O OH O OH O OH O OH O O O O HO O HO HO O

(CH2 ) * 1 or 3 (CH ) * (CH2 ) * (CH2 ) (CH ) * 2 1,2 or 3 1,2 or 3 1,2 or 3 2 1,2 or 3

Isobranched Aliphatic Chains 705, 691 and 677 960, 946 and 932 736, 722 and 708 584, 570 and 556 662 and 634 m/z negative ions m/z negative ions m/z negative ions m/z positive ions m/z negative ions

Figure 1. Chemical structures of P. gingivalis complex lipids are given with corresponding mass to charge ratios (m/z). PGDHC consists of a fatty acid, 3-OH isoC17:O, and a long chain base that varies from 17-19 carbons in length. When PGDHC is treated with NaOCH3, resultant de-esterification yields unsubstituted PGDHC (unsub PGDHC) which is essentially structurally identical to PGDHC minus a 15-carbon isobranched methyl ester. Both PGDHC and Unsub PGDHC contain a phosphoglycerol head group. Free DHC is structurally identical to unsub PGDHC minus the phosphoglycerol head group. Two other previously investigated P. gingivalis lipids, phosphoethanolamine and phosphatidylethanolomine, were not tested in this experiment, but were included in this figure for completeness. Figure was provided by Frank Nichols.

33

Figure 2. ESI – spectrum of total lipid extract of P. gingivalis. Predominant ion abundances (m/z: 960, 946, 932) correspond to PGDHC. Lower spectrum corresponds to solvent.

34

Figure 3. ESI - spectrum of PGDHC fractions after separation of P. gingivalis total lipid extract with dHPLC. Purity of sample can be appreciated by comparing relative ion abundances between PGDHC (m/z: 960, 946, 932) seen in upper spectrum and solvent in lower spectrum.

35

Figure 4. ESI - spectrum of PGDHC – C15:0 fractions after separation of P. gingivalis total lipid extract with dHPLC. Again, purity of sample can be determined by comparison of relative ion abundance between PGDHC-C15:0 and other signals including solvent spectrum below. High ion abundance signal to the left in the spectrum corresponds to negligible fatty acid moieties.

36

Figure 5. UV-vis spectrum of rHPLC at 205 nm. Second peak corresponds to fDHC determined by ESI + mass spectrometry (not shown).

37

Figure 6. UV-vis spectrum monitored at 205 nm of fDHC fraction collected from first separation of P. gingivalis. Free DHC fraction corresponds with strongest peak determined by ESI + mass spectrometry (shown in Figure 7).

38

Figure 7. ESI + spectrum of P. gingivalis fDHC fractions after separation P. gingivalis total lipid extract with rHPLC. Purity of sample can be appreciated by comparing relative ion abundances between fDHC (m/z: 584, 570, 556) seen in upper spectrum and solvent in lower spectrum.

39

Figure 8. ESI + spectrum of synthetic fDHC fraction (A). Purity of sample can be appreciated by comparing relative ion abundances between synthetic fDHC (A) with an isobranched C17:0 long chain base (m/z: 556) seen in upper spectrum and solvent in lower spectrum.

40

Figure 9. NMR spectra of P. gingivalis fDHC fractions (red and blue spectra) and synthetic fDHC (black spectrum). A high degree of similarity between bacterial and synthetic fDHC can be appreciated.

41

Panel A Panel B

Panel C Panel D

Figure 10. Phase contrast photomicrographs at 100x magnification of HGF in 35mm culture wells with ethanol vehicle (Panel A), total lipid extract (Panel B), PGDHC (Panel C), and fDHC (Panel D) for 24 hours. Normal HGF morphology is noted for ethanol vehicle and fDHC (Panels A and D). Cellular apoptosis is noted for Total lipid extract and PGDHC (Panels B and C).

42

1000 n=3 PGE 2 PGF n=3 n=3 2 α 800 Prostaglandin Release 600 n=3 (ng/3ml) 400

200

0 EtOH PG (20 ug) DHC P. g. FDHC ( 20 ug) P. FDHC g. Synthetic (20 ug) FDHC

Fibroblast Treatment

Figure 11. Comparison of HGF production of PGE2 and PGF2α . Error bars represent standard deviations for each sample.

43

2

n=3

n=3 n=3 n=3

PGE (T/C) 2 1

0 PG (20 ug) DHC P. g. FDHC ( 20 ug) P. FDHC g. Unsub PG(20 ug) DHC Synthetic (20 ug) FDHC

Fibroblast Treatment

Figure 12. Prostaglandin release given as treatment/control (T/C) for each sample examined. Ethanol control is not shown. Error bars represent standard deviations.

44

3

n=11 n=6 2 * n=3 * PGE (T/C) 2

1

0 PG (20 ug) DHC P. g. FDHC ( 20 ug) P. FDHC g. Synthetic (20 ug) FDHC * p< 0.05 by Fisher PLSD Fibroblast Treatment

Figure 13. Prostaglandin release given as treatment/control (T/C) for each sample examined. Ethanol control is not shown. Error bars represent standard deviations.

45

3

n=4

2 n=3 n=4 PGE (T/C) n=4 n=4 2

1

0 PG (20 ug) DHC P. g. FDHC ( 20 ug) P. FDHC g. P. g. FDHC ( 0.2 ug) P. FDHC g. ( 2.0 ug) P. FDHC g. Synthetic (20 ug) FDHC

Fibroblast Treatment

Figure 14. Prostaglandin release given as treatment/control (T/C) for each sample examined. Ethanol control is not shown. Error bars represent standard deviations.

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