University of Iowa
Iowa Research Online
Theses and Dissertations
Spring 2013
Oral mucosal lipids are antimicrobial against
Porphyromonas gingivalis, induce ultrastructural
damage, and alter bacterial lipid and protein
compositions
Carol Lea Fischer
University of Iowa
Copyright 2013 Carol Lea Fischer
This dissertation is available at Iowa Research Online: http://ir.uiowa.edu/etd/2494
Recommended Citation
Fischer, Carol Lea. "Oral mucosal lipids are antimicrobial against Porphyromonas gingivalis, induce ultrastructural damage, and alter
bacterial lipid and protein compositions." PhD (Doctor of Philosophy) thesis, University of Iowa, 2013.
http://ir.uiowa.edu/etd/2494.
Follow this and additional works at: http://ir.uiowa.edu/etd
Part of the Oral Biology and Oral Pathology Commons
ORAL MUCOSAL LIPIDS ARE ANTIBACTERIAL AGAINST PORPHYROMONAS
GINGIVALIS, INDUCE ULTRASTRUCTURAL DAMAGE, AND ALTER
BACTERIAL LIPID AND PROTEIN COMPOSITIONS
by
Carol Lea Fischer
An Abstract
Of a thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Oral Science
in the Graduate College of
The University of Iowa
May 2013
Thesis Supervisor: Professor Kim A. Brogden
ABSTRACT
Periodontal disease is a chronic inflammation of the gingiva and periodontium
that leads to progressive destruction and irreversible damage to the supportive structures
of the teeth. It affects nearly half of the United States population and is a particular risk
factor in adults older than 65 years of age. Oral microorganisms assemble in plaque as a
polymicrobial biofilm and Porphyromonas gingivalis, an important secondary colonizer
in oral biofilms, has been implicated in periodontal disease. Although the protective
functions of various salivary molecules such as antimicrobial proteins have been
delineated, lipids present in saliva and on the oral mucosa have been largely ignored and
there is growing evidence that the role of lipids in innate immunity is more important
than previously realized. In fact, recent studies suggest that sphingoid bases and fatty
acids, which exhibit potent broad spectrum antimicrobial activity against a variety of
bacteria and fungi, are likely important innate immune molecules involved in the defense
against oral bacterial and fungal infections. However little is known about their spectrum
of activity or mechanisms of action. In addition, the effects of these lipids that are
endogenous to the oral cavity have not been explored against oral bacteria. In this study I
hypothesized that oral mucosal and salivary lipids exhibit dose-dependent antimicrobial
activity against P. gingivalis and alter cell morphology and metabolic events. To test this
hypothesis, I first examined the effects of two fatty acids: sapienic acid and lauric acid,
and three sphingoid bases: sphingosine, dihydrosphingosine, and phytosphingosine,
against a variety of gram-positive and gram-negative bacteria including P. gingivalis.
Using broth microdilution assays to determine minimum inhibitory and minimum
bactericidal concentrations, I show that antimicrobial activity against bacteria is dosedependent, lipid specific, and microorganism specific. Kill kinetics were also variable
across each bacteria-lipid combination. Upon examination of select bacteria-lipid
combinations via scanning and transmission electron microscopy, different morphologies
were evident across all treatments, demonstrating differential activity of each lipid for a
particular bacterium as well as for each bacterium across different lipids. In addition, all
sphingoid bases and fatty acids were taken up and retained in association with P.
gingivalis cells and could be extracted along with bacterial lipids and separated using thin
layer chromatography. Using a combination of two-dimensional in-gel electrophoresis
and Western blots followed by mass spectroscopy and n-terminus degradation
sequencing, I show that sapienic-acid treatment induces a unique stress response in P.
gingivalis, as evidenced by the ability of P. gingivalis to upregulate a set of proteins
involved in fatty acid biosynthesis metabolism and energy production, protein processing,
cell adhesion, and virulence. Finally, utilizing flow cytometry and confocal microscopy,
I assessed the effects of oral antimicrobial lipids against a representative host cell and
describe oral lipid concentrations that are both antimicrobial to P. gingivalis cells and
non-cytotoxic to the representative host cells tested. Combined, these data strongly
suggest that sphingoid bases and fatty acids found within the saliva and on oral mucosa
likely do contribute to the innate antimicrobial activity of saliva, mucosal surfaces, and
skin and this dose-dependent activity is both lipid specific and bacteria specific. This
information adds to current knowledge of the innate functions of endogenous lipids in the
oral cavity. With bacterial resistance to current antibiotics increasing, the exploration of
new antimicrobial agents is important and these lipid treatments may be beneficial for
prophylactic treatments or therapeutic intervention of infection by supplementing the
natural immune function of endogenous lipids on skin and other mucosal membranes.
Abstract Approved: ____________________________________
Thesis Supervisor
____________________________________
Title and Department
____________________________________
Date
ORAL MUCOSAL LIPIDS ARE ANTIBACTERIAL AGAINST PORPHYROMONAS
GINGIVALIS, INDUCE ULTRASTRUCTURAL DAMAGE, AND ALTER
BACTERIAL LIPID AND PROTEIN COMPOSITIONS
by
Carol Lea Fischer
A thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Oral Science
in the Graduate College of
The University of Iowa
May 2013
Thesis Supervisor: Professor Kim A. Brogden
Copyright by
CAROL LEA FISCHER
2013
All Rights Reserved
Graduate College
The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
_______________________
PH.D. THESIS
_______________
This is to certify that the Ph.D. thesis of
Carol Lea Fischer
has been approved by the Examining Committee
for the thesis requirement for the Doctor of Philosophy
degree in Oral Science at the May 2013 graduation.
Thesis Committee: ___________________________________
Kim A. Brogden, Thesis Supervisor
___________________________________
Philip Wertz
___________________________________
David Drake
___________________________________
Georgia Johnson
___________________________________
Mary Wilson
To Benjamin and Samuel, my reasons for living: this document is proof that with
God’s help you can accomplish anything your heart desires. Do not ever let anyone tell
you otherwise. This is the tribute I wish to leave with you.
To Scott, my husband and my best friend: thank you for being my wall, for letting
me lean on you when things got rough, for all your support, encouragement, patience,
dedication to my dreams, and for loving me even when I was too busy to stop and remind
you of how very thankful I am that you’re here.
Words cannot express how much I love each of you.
ii
ACKNOWLEDGMENTS
I would like to express my deep appreciation and gratitude to Dr. Kim Brogden,
my advisor and friend, for his extraordinary mentorship, support, and patient guidance
throughout this entire process – from application to completion. The combination of Dr.
Brogden’s intellect and genuinely good nature made my time here a pleasure and I am
truly fortunate to have had the opportunity to work with him. This thesis would not have
been possible without the support, guidance, and thought-provoking questions of Dr.
Brogden and the rest of my committee members. Dr. Philip Wertz was a veritable
encyclopedia of lipid knowledge upon which I drew continuously. Dr. Mary Wilson, Dr.
David Drake, and Dr. Georgia Johnson, in addition to giving of their time to serve on my
committee, provided a steady source of support, advice, and encouragement.
Collectively, my committee provided an atmosphere that fostered independent and
critical thinking, learning, and personal growth and I will be forever grateful to each and
every one of them.
I am extremely appreciative of several funding sources supporting my research
including NIH/NIDCR T32 DE014678, NIH/NIDCR R01 DE018032 (Dr. Wertz’ R01),
and NIH/NIDCR R01 DE014390 (Dr. Brogden’s R01). I also want to thank Dr. Leslie
Mehalick for the use of one of her figures and data. I am indebted to the College of
Dentistry, my lab members, and all my friends here, for their continuous moral support
and encouragement, for listening to me talk about my research, and for providing a
comfortable family atmosphere that contributed to a positive learning environment.
Many thanks also to Dr. Deborah Dawson and Derek Blanchette for all their statistical
expertise and for answering a million questions. I would also like to thank my graduate
student and post-grad friends for their support, feedback, and friendship.
Several of my undergraduate professors, Dr. Kenneth Andrews, Dr. Michael Bay,
Dr. Charlie Biles, and Dr. Terry Cluck, who contributed to my love of science and
iii
encouraged me to pursue a doctorate, also deserve my gratitude for helping me find my
way here and showing support and encouragement as I pursued this degree. I would also
like to thank my large group of family and friends for their love and support, for
accepting nothing less than completion from me, and for being there when I needed
encouragement or a nudge in the right direction. Specifically I want to thank Benjamin
and Samuel for their willingness to share me with “school” for the past eleven years
while I completed both my undergraduate and graduate degrees. Their understanding,
support, encouragement, and unfailing love kept me going when things were difficult.
And last, but certainly not least, a special thanks to my husband, Scott for being my solid
wall of strength and for shouldering more than his fair share of the household duties
during comps and the writing of this dissertation. Most importantly, I am thankful for my
faith in God, who is the ultimate provider of all my strength and everything good in my
life.
iv
ABSTRACT
Periodontal disease is a chronic inflammation of the gingiva and periodontium
that leads to progressive destruction and irreversible damage to the supportive structures
of the teeth. It affects nearly half of the United States population and is a particular risk
factor in adults older than 65 years of age. Oral microorganisms assemble in plaque as a
polymicrobial biofilm and Porphyromonas gingivalis, an important secondary colonizer
in oral biofilms, has been implicated in periodontal disease. Although the protective
functions of various salivary molecules such as antimicrobial proteins have been
delineated, lipids present in saliva and on the oral mucosa have been largely ignored and
there is growing evidence that the role of lipids in innate immunity is more important
than previously realized. In fact, recent studies suggest that sphingoid bases and fatty
acids, which exhibit potent broad spectrum antimicrobial activity against a variety of
bacteria and fungi, are likely important innate immune molecules involved in the defense
against oral bacterial and fungal infections. However little is known about their spectrum
of activity or mechanisms of action. In addition, the effects of these lipids that are
endogenous to the oral cavity have not been explored against oral bacteria. In this study I
hypothesized that oral mucosal and salivary lipids exhibit dose-dependent antimicrobial
activity against P. gingivalis and alter cell morphology and metabolic events. To test this
hypothesis, I first examined the effects of two fatty acids: sapienic acid and lauric acid,
and three sphingoid bases: sphingosine, dihydrosphingosine, and phytosphingosine,
against a variety of gram-positive and gram-negative bacteria including P. gingivalis.
Using broth microdilution assays to determine minimum inhibitory and minimum
bactericidal concentrations, I show that antimicrobial activity against bacteria is dosedependent, lipid specific, and microorganism specific. Kill kinetics were also variable
across each bacteria-lipid combination. Upon examination of select bacteria-lipid
combinations via scanning and transmission electron microscopy, different morphologies
v
were evident across all treatments, demonstrating differential activity of each lipid for a
particular bacterium as well as for each bacterium across different lipids. In addition, all
sphingoid bases and fatty acids were taken up and retained in association with P.
gingivalis cells and could be extracted along with bacterial lipids and separated using thin
layer chromatography. Using a combination of two-dimensional in-gel electrophoresis
and Western blots followed by mass spectroscopy and n-terminus degradation
sequencing, I show that sapienic-acid treatment induces a unique stress response in P.
gingivalis, as evidenced by the ability of P. gingivalis to upregulate a set of proteins
involved in fatty acid biosynthesis metabolism and energy production, protein processing,
cell adhesion, and virulence. Finally, utilizing flow cytometry and confocal microscopy,
I assessed the effects of oral antimicrobial lipids against a representative host cell and
describe oral lipid concentrations that are both antimicrobial to P. gingivalis cells and
non-cytotoxic to the representative host cells tested. Combined, these data strongly
suggest that sphingoid bases and fatty acids found within the saliva and on oral mucosa
likely do contribute to the innate antimicrobial activity of saliva, mucosal surfaces, and
skin and this dose-dependent activity is both lipid specific and bacteria specific. This
information adds to current knowledge of the innate functions of endogenous lipids in the
oral cavity. With bacterial resistance to current antibiotics increasing, the exploration of
new antimicrobial agents is important and these lipid treatments may be beneficial for
prophylactic treatments or therapeutic intervention of infection by supplementing the
natural immune function of endogenous lipids on skin and other mucosal membranes.
vi
TABLE OF CONTENTS
CHAPTER 1 INTRODUCTION .........................................................................................1
Microbiota of the Oral Cavity...........................................................................2
Pathogens of Periodontal Disease .....................................................................3
Porphyromonas gingivalis and Periodontal Disease ........................................4
Host Immunity to Oral Bacterial Infections .....................................................5
Oral Lipids as Host Innate Immune Factors .....................................................6
Epithelial lipids of the oral cavity .............................................................7
Sphingoid base structure and function ...............................................9
Sebaceous lipids of the oral cavity ..........................................................10
Fatty acid structure and function ......................................................11
Research Aims ................................................................................................12
CHAPTER 2 ANTIBACTERIAL ACTIVITY OF SPHINGOID BASES AND
FATTY ACIDS AGAINST GRAM-POSITIVE AND GRAMNEGATIVE BACTERIA ...............................................................................14
Materials and Methods ...................................................................................15
Bacterial species and growth conditions .................................................15
Preparation of lipids ................................................................................16
Antimicrobial assays ...............................................................................16
Killing kinetics assays .............................................................................17
Statistical analyses ...................................................................................18
Results.............................................................................................................18
Discussion .......................................................................................................20
CHAPTER 3 SPHINGOID BASES INDUCE ULTRASTRUCTURAL DAMAGE
IN ESCHERICHIA COLI AND STAPHYLOCOCCUS AUREUS AND
ALTER THEIR LIPID COMPOSITION .......................................................32
Materials and Methods ...................................................................................34
Bacterial species and growth conditions .................................................34
Preparation of lipids ................................................................................34
Preparation of lipid-damaged bacterial cells ...........................................34
Scanning electron microscopy .................................................................34
Transmission electron microscopy ..........................................................35
Cell dimensions and statistical analysis ..................................................35
Isolation of lipids from bacteria ..............................................................36
Lipid analysis...........................................................................................37
Results.............................................................................................................37
Scanning electron microscopy .................................................................37
Transmission electron microscopy ..........................................................39
Thin layer chromatography .....................................................................40
Discussion .......................................................................................................40
CHAPTER 4 ORAL MUCOSAL LIPIDS ARE ANTIBACTERIAL AGAINST
PORPHYROMONAS GINGIVALIS, INDUCE ULTRASTRUCTURAL
DAMAGE, AND ALTER BACTERIAL LIPID AND PROTEIN
COMPOSITIONS ...........................................................................................50
Materials and Methods ...................................................................................52
Bacterial species and growth conditions .................................................52
Preparation of lipids ................................................................................52
Antimicrobial assay .................................................................................52
vii
Kill kinetics .............................................................................................53
Ultrastructural analyses of lipid-exposed bacterial cells .........................53
Lipid analysis...........................................................................................55
Protein analyses .......................................................................................56
Statistical analyses ...................................................................................57
Results.............................................................................................................58
Discussion .......................................................................................................61
CHAPTER 5 ORAL MUCOSAL LIPID CYTOTOXICITY: A STUDY USING
DENDRITIC CELLS ......................................................................................79
Materials and Methods ...................................................................................81
Preparation of lipids ................................................................................81
Preparation of dendritic cells ...................................................................81
Flow cytometry ........................................................................................82
Confocal microscopy ...............................................................................83
Results.............................................................................................................83
Discussion .......................................................................................................86
CHAPTER 6 CONCLUSIONS .......................................................................................112
Potential Mechanisms of Activity ................................................................112
Implications for Prophylactic or Therapeutic Treatments ............................114
APPENDIX ......................................................................................................................115
REFERENCES ................................................................................................................126
viii
LIST OF TABLES
Table 2.1.
Inhibitory and bactericidal activity of sphingoid bases and fatty acids
for gram-negative and gram-positive bacteria. .............................................23
Table 2.2.
Time to zero comparisons of lipid kill kinetics for gram-positive and
gram negative bacteria. .................................................................................26
Table 2.3.
Trapezoidal AUC comparisons of lipid treatments as a summary
measure of bacterial viability over the treatment time course. .....................27
Table 3.1.
Visual surface area descriptive statistics for untreated and sphingoid
base-treated E. coli and S. aureus. ................................................................44
Table 3.2.
E. coli pairwise exact Wilcoxon Rank Sum treatment comparisons of
the visual surface area. ..................................................................................45
Table 3.3.
S. aureus pairwise exact Wilcoxon Rank Sum treatment comparisons
of the visual surface area...............................................................................46
Table 4.1.
Minimum lipid concentrations required to inhibit or kill P. gingivalis. .......67
Table 4.2.
Pairwise comparisons of the time required to kill P. gingivalis by each
of the lipid treatments. ..................................................................................68
Table 4.3.
AUC analysis of kill kinetics. .......................................................................69
Table 4.4.
Identification of P. gingivalis upregulated proteins upon treatment
with sapienic acid. .........................................................................................70
Table 5.1.
Antimicrobial and cytotoxic activity of sphingoid bases..............................90
Table A.1.
P. gingivalis stress responses. ....................................................................116
ix
LIST OF FIGURES
Figure 2.1.
Kill kinetics of lipid treatments against gram-positive and gramnegative bacteria. .........................................................................................29
Figure 3.1.
SEM images of E. coli and S. aureus untreated and treated with
sphingoid bases............................................................................................47
Figure 3.2.
TEM images of E. coli and S. aureus untreated and treated with
sphingoid bases............................................................................................48
Figure 3.3.
Association of E. coli and S. aureus lipids with sphingoid bases after
treatment ......................................................................................................49
Figure 4.1.
Kill kinetics for all lipid treatments against P. gingivalis. ..........................73
Figure 4.2.
SEM micrographs showing the effects of sphingoid base and fatty
acid treatment on P. gingivalis. ...................................................................74
Figure 4.3.
TEM micrographs showing the effects of sphingoid base and fatty
acid treatment on P. gingivalis. ...................................................................75
Figure 4.4.
Association of antimicrobial lipids with P. gingivalis lipids after
treatment. .....................................................................................................76
Figure 4.5.
SDS-PAGE separation of proteins in untreated and sapienic acidtreated P. gingivalis .....................................................................................77
Figure 4.6.
2D-DIGE gel showing P. gingivalis protein differences in untreated
and sapienic acid-treated samples ...............................................................78
Figure 5.1.
Cytotoxicity of sphingoid bases against DCs ..............................................91
Figure 5.2.
Confocal micrographs of untreated DCs. ....................................................92
Figure 5.3.
Confocal micrographs of DCs treated with 80 µM sphingosine .................93
Figure 5.4.
Confocal micrographs of killed DCs ...........................................................94
Figure 5.5.
Confocal micrographs of DCs treated with 80 µM
dihydrosphingosine .....................................................................................95
Figure 5.6.
Confocal micrographs of DCs treated with 80 µM phytosphingosine ........96
Figure 5.7.
Confocal micrographs of DCs treated with 80 µM glycerol
monolaurate .................................................................................................97
Figure 5.8.
Confocal micrographs showing sphingoid base affinity for C12resazurin dye ................................................................................................98
Figure 5.9.
Flow cytometry data showing sphingoid base affinity for C12resazurin dye..............................................................................................100
x
Figure 5.10. Flow cytometric analyses of sphingoid base cytotoxicity against DCs. ...101
Figure 5.11. Cytotoxicity of sphingoid bases for DCs...................................................106
Figure 5.12. Forward/side scatter graphs of flow cytometric data for treatment of
DCs with phytosphingosine (Phy) at 80 µM concentrations.....................107
Figure 5.13. Confocal micrographs of DCs treated with 5 µM sphingosine .................108
Figure 5.14. Confocal micrographs of DCs treated with 5 µM dihydrosphingosine.....109
Figure 5.15. Confocal micrographs of DCs treated with 5 µM phytosphingosine ........110
Figure 5.16. Confocal micrographs of DCs treated with 5 µM glycerol
monolaurate. ..............................................................................................111
xi
1
CHAPTER 1
INTRODUCTION
Infection and inflammation in the oral cavity ranges from gingivitis, a mild and
reversible inflammation of the gingiva, to aggressive periodontitis, a chronic
inflammation of the gingiva and periodontium that leads to progressive destruction of the
periodontal ligament and alveolar bone. Periodontitis results in the formation of
periodontal pockets below the gingiva and adjacent to the tooth surface which become
heavily colonized with bacteria, leading to chronic inflammation of the supporting tissues
of the teeth and subsequent loss of connective tissue and bone (Darveau 2010). One
theory is that the host mounts an exaggerated immune response that is not only
ineffectual but contributes to tissue damage and the progression of disease via host
proteinases (Berglundh and Donati 2005). Damage caused by periodontal disease (e.g.
gingival detachment and bone loss) is irreversible but treatment can halt the progression
of disease.
Periodontitis occurs in just over 47% of the population of the United States with a
prevalence of 8.7, 30.0, and 8.5% for mild, moderate, and severe periodontitis,
respectively (Eke et al. 2012) and is dependent upon oral hygiene, socio-economic status,
and other environmental, genetic and metabolic risk factors that contribute to host
susceptibility. Examples of factors linked to increased risk of periodontal disease include
tobacco use, alcohol use, diabetes, and stress (Pihlstrom et al. 2005). If untreated,
periodontal disease not only affects oral health, but potentially systemic health. Several
studies describe evidence linking pathogenic periodontal microorganisms to systemic
diseases such as cardiovascular and pulmonary diseases (Beck and Offenbacher 2005,
Joshipura et al. 2003, Offenbacher et al. 1998) as well as preterm births (Offenbacher et
al. 1998).
2
Microbiota of the Oral Cavity
The oral cavity of healthy individuals contains a menagerie of bacterial, viral,
fungal, and protozoan species colonizing both hard and soft tissue surfaces which make
up several distinct microbial habitats (Dewhirst et al. 2010, Wade 2012). The human
mouth, second to only the colon in species diversity (Wade 2012), harbors billions of
bacteria representing an estimated 700 – 1000 phylotypes, less than half of which are
cultivatable using standard microbiological methods (Aas et al. 2005, Dewhirst et al.
2010, Socransky et al. 1998, Wade 2012). Not all of these bacteria are present in an
individual simultaneously. It is estimated that individuals harbor approximately 100 –
200 species in the oral cavity at any given time (Siqueira and Rocas 2010) and bacterial
community profiles differ depending upon the surface of colonization (e.g. different oral
structures and tissues) (Aas et al. 2005, Dewhirst et al. 2010).
The Human Oral Microbiome Database (HOMD; www.homd.org) describes 662
oral bacterial phylotypes from 13 phyla and provides comprehensive information such as
genome and 16S rRNA (Dewhirst et al. 2010). Six of the 13 phyla (Firmicutes,
Bacteroidetes, Proteobacteria, Actinobacteria, Spirochaetes, and Fusobacteria) contain
96% of the known oral bacterial phylotypes while the remaining phyla (Euryarchaeota,
Chlamydia, Chloroflexi, Synergistetes, Tenericutes, SR1, and TM7) contain 4% of the
known oral phylotypes (Dewhirst et al. 2010). Streptococcus and Actinomyces represent
the most abundant genera while Prevotella, of the phylum Bacteroidetes, is the largest
genus, containing approximately 50 species (Dewhirst et al. 2010, Kolenbrander et al.
2010, Siqueira and Rocas 2010, Wade 2012).
The majority of oral bacteria are associated with, or at least compatible with,
periodontal health (Haffajee et al. 1998, Wade 2012). These health-associated bacteria
primarily include genera from five phyla including Firmucutes, Actinobacteria,
Proteobacteria, Bacteroidetes, and Fusobacteria (Aas et al. 2005, Zaura et al. 2009) and
their presence likely helps prevent colonization by pathogenic bacteria (Vollaard and
3
Clasener 1994). Periodontally healthy individuals’ bacterial load is typically low (102 –
103 bacteria/gram of plaque) and comprises primarily the gram-positive bacteria
Streptococcus or Actinomyces (Darveau 2010); however, low numbers of diseaseassociated bacterial species often exist in the absence of disease. Poor oral hygiene and
other host or environmental cues may confer selective growth advantage to pathogenic
microorganisms, resulting in a population shift from largely health-associated
microorganisms to a more disease-associated microbiome and an increase in total
bacterial load (Amano 2010, Kolenbrander et al. 2010, Lamont and Jenkinson 2000,
Pennisi 2005, Socransky and Haffajee 1992).
Pathogens of Periodontal Disease
Although no single etiologic agent has been identified in the development of
periodontitis, specific genera, including Porphyromonas (P. gingivalis, P. endodontalis),
Treponema (T. denticola, T. socranskii), Tannerella forsythia, Prevotella (P. intermedia,
P. nicrescens, P. baroniae), Aggregatibacter actinomycetemcomitans, Fusobacterium
nucleatum, Filifactor alocis, Eubacterium (E. nodatum, E. sulci), Parvimonas micra, and
others are strongly associated with periodontal inflammation and related diseases
(Haffajee et al. 1998, Ledder et al. 2007, Siqueira and Rocas 2010, Socransky et al. 1998,
Wade 2012). Periodontal disease is typically associated with a shift from a largely grampositive community (e.g. streptococci and actinomycetes) to a largely gram-negative
community characterized by higher numbers of putative periodontal pathogens and
leading to an increase in total microbial load (Ledder et al. 2007). As with many
diseases, periodontitis is influenced by a consortium of microorganisms, known as
biofilms, rather than by single pathogens (Dewhirst et al. 2010, Kolenbrander et al. 2010)
and it is likely that within these microbial communities, interactions between commensal
and pathogenic bacteria also contribute to the pathogenicity of the oral microbial
community (Hajishengallis et al. 2011, Offenbacher 1996).
4
Porphyromonas gingivalis and Periodontal Disease
P. gingivalis is a gram-negative, non-motile, asaccharolytic, strictly anaerobic
coccobacillus that is consistently associated with periodontitis (Holt et al. 1999, Hutter et
al. 2003, Ledder et al. 2007, Paster et al. 2001, Socransky and Haffajee 1992, Socransky
et al. 1998). P. gingivalis is more likely to be present in patients with periodontitis and
shows a strong positive relationship with diagnostic parameters for periodontitis,
including gingival recession, increased sulcular pocket depth and bleeding upon probing
(Hutter et al. 2003, Socransky and Haffajee 1992, Socransky et al. 1998). In addition,
Hajishengallis and colleagues recently demonstrated that although P. gingivalis does not
independently cause periodontal disease in a germ-free murine model, low numbers of P.
gingivalis can disrupt host homeostasis through actions requiring both commensal
microorganisms and complement, leading to inflammation and periodontal disease
(Hajishengallis et al. 2011).
P. gingivalis produces multiple virulence factors that allow successful
colonization and support evasion of host defenses, many of which contribute to
inflammation and destruction of host tissue (Holt et al. 1999). Adhesin molecules (e.g.
fimbraie and hemagglutinins) promote attachment (Holt et al. 1999, Lamont and
Jenkinson 1998, Lamont and Jenkinson 2000, Offenbacher 1996) while proteolytic
enzymes (e.g. cysteine proteinases and hemagglutinins) are capable of degrading multiple
substrates in the gingival crevice, facilitating nutrient acquisition and contributing to host
tissue degradation (Holt et al. 1999, Lamont and Jenkinson 1998, Lamont and Jenkinson
2000).
A number of virulence factors produced by P. gingivalis are capable of
modulating host immune processes. P. gingivalis and its toxic by-products can activate
the complement system (Lamont and Jenkinson 1998), induce neutrophil chemotactic
factors (Holt et al. 1999), and induce expression of both anti-inflammatory and
proinflammatory cytokines and chemokines (Lamont and Jenkinson 1998). In addition,
5
P. gingivalis produces several immunoglobulin proteinases and enzymes capable of
degrading complement proteins, cytokines, antimicrobial proteins, and neutrophil
receptors for immunoglobulin and complement (Holt et al. 1999, Lamont and Jenkinson
1998, Lamont and Jenkinson 2000). The arginine-specific cysteine proteinase, arggingipain, is a good example of a proteinase that functions in several ways to alter host
defense mechanisms. In addition to serving as a potent chemotactic molecule for
neutrophils, arg-gingipains can cleave immunoglobulins and select complement proteins,
and cleave neutrophil receptors for complement and immunoglobulins (Holt et al. 1999).
P. gingivalis can also evade phagocytosis through secretion of a polysaccharide capsule
which prevents opsonization by complement and immunoglobulins (Holt et al. 1999,
Lamont and Jenkinson 1998). In addition, P. gingivalis can form surface blebs (outer
membrane vesicles) which bind complement and immunoglobulins before they can reach
the bacterial cells (Holt et al. 1999, Lamont and Jenkinson 1998).
Host Immunity to Oral Bacterial Infections
Both the adaptive and innate defense mechanisms play a role in periodontal
disease; however, the relative contribution of each in the clearance of disease versus
contribution to disease is still incompletely defined (Lamont and Jenkinson 1998). In
healthy oral tissue, neutrophil-mediated phagocytosis plays a major role in controlling
overgrowth of periodontal bacteria (Darveau 2010, Gemmell et al. 2007, Offenbacher
1996). Approximately 30,000 polymorphonuclear leukocytes (PMN) – mainly
neutrophils – transit through the periodontal tissue every minute, forming a barrier
between the host tissue and dental plaque (Darveau 2010). In healthy individuals,
activation of complement through the classical pathway leads to opsonization and
clearance of periodontal pathogens by neutrophil-mediated phagocytosis (Offenbacher
1996). In addition, a diverse array of specific and non-specific innate immune factors
present in saliva and on mucosal surfaces help maintain periodontal health. More than 45
6
antimicrobial proteins (AMP) are grouped into functional families that include cationic
peptides, metal ion chelators, protease inhibitors, peroxidases, bacterial adhesins and
agglutinators, and enzymes directed at the bacterial cell wall (Gorr 2009, Gorr 2012).
AMPs and other salivary molecules act on bacteria by a variety of mechanisms including
direct antimicrobial activity (e.g. defensins, histatins, cystatins, lactoferrin, sphingoid
bases, fatty acids), disruption of bacterial adhesion and co-adhesion with other
microorganisms (e.g. histatins, lysozyme, fibronectin), agglutinins (e.g. mucins, secretory
IgA, lysozyme, fibronectin), and inactivation of bacterial proteases (e.g. histatins,
cystatins, secretory leukoprotease inhibitor) (Bibel et al. 1993, Bratt et al. 2011, Brogden
et al. 2011, Drake et al. 2008, Fischer et al. 2012, Gorr 2009, Gorr 2012, Lamont and
Jenkinson 1998, Lamont and Jenkinson 2000, Tenovuo et al. 1987).
Specific triggers for the switch from periodontal health to disease are still elusive
but as previously discussed, the answer is likely complicated, involving a combination of
oral hygiene and other environmental, genetic, and metabolic risk factors. Additionally,
as discussed in the previous section, P. gingivalis has the ability to disrupt many innate
host defense mechanisms through a variety of proteinases and other virulence factors.
For example, P. gingivalis is able to disrupt nearly every aspect of neutrophil recruitment
and activity including inhibition of neutrophil chemotaxis, degradation of complement
and immunoglobulins, cleavage of complement receptors from neutrophils, and secretion
of a capsule (Gemmell et al. 2007, Holt et al. 1999, Lamont and Jenkinson 1998, Lamont
and Jenkinson 2000).
Oral Lipids as Host Innate Immune Factors
Although less well known, sphingoid bases and certain fatty acids found on the
surface of the oral mucosa and in saliva exhibit antimicrobial activity against a variety of
gram-positive and gram-negative bacteria. Oral antimicrobial lipids are produced by
either oral epithelium (e.g. sphingoid bases) or sebaceous follicles (e.g. fatty acids). Both
7
sphingoid bases and fatty acids are also present in saliva. In the following sections I will
discuss the production and secretion of the antimicrobial lipids of epithelial and
sebaceous origin along with structure and function.
Epithelial lipids of the oral cavity
The production and secretion of lipids in the oral cavity is primarily the function
of the epithelia and sebaceous glands. The hard palate and gingiva are covered by
stratum corneum consisting of flattened, keratin-filled cells embedded in a lipid matrix;
the buccal region, underside of the tongue, and floor of the mouth do not have a stratum
corneum but the outer one-third of these non-keratinized epithelial regions consists of
metabolically inactive cells which provide a permeability barrier similar to stratum
corneum of keratinized epithelia (Downing et al. 1993, Law et al. 1995b).
Lipids are synthesized in the viable portion of the epithelium; therefore, all the
epithelium in the oral cavity produces lipids, including phospholipids (e.g.
sphingomyelin, phosphatidylethanolamine, phosphatidylserine, phosphatidylinositol),
glycosylceramides, ceramides, sterols, and sterol esters. In keratinizing epithelia, these
lipids are packaged in lamellar granules along with a variety of hydrolytic enzymes (e.g.
ceramidase, sphingomyelinase) and pushed outward toward the stratum corneum where
the lipids are extruded into the intercellular spaces between the stratum granulosum and
stratum corneum (Brogden et al. 2011, Downing et al. 1993, Drake et al. 2008). In nonkeratinizing epithelia, lipids are packaged into a similar small secretory organelle, which
extrudes its contents into the intercellular space at the location where the permeability
barrier forms (Law et al. 1995a). Lipids in stratum corneum of keratinizing epithelia and
the outer one-third of the non-keratinizing epithelia include phospholipids,
glucosylceramides, ceramides, fatty acids, cholesterol, and cholesterol esters but there are
higher proportions of phospholipids and glycosylceramides in the barrier layers of non-
8
keratinizing epithelia (Law et al. 1995a). Phospholipids, glucosylceramides, and
phosphoglycerides are then hydrolyzed to produce ceramides and fatty acids.
Ceramides, consisting of fatty acids and α-hydroxyacids or ω-hydroxyacids
amide-linked to a sphingoid base (sphingosine, dihydrosphingosine, phytosphingosine, or
6-hydroxyspingosine), are the source of sphingoid bases (Wertz and Downing 1990a).
Free sphingoid bases and long-chain fatty acids are liberated from ceramides via the
action of ceramidases found both in the viable portion of the epidermis and in the stratum
corneum (Law et al. 1995b). Worth noting here is that fatty acids of epithelial origin that
reach the skin surface are primarily long straight-chain fatty acids (20 – 28 carbons) and
have no demonstrated antimicrobial activity (Brogden et al. 2011, Wertz et al. 1987);
therefore, they should not be confused with the antimicrobial fatty acids of sebaceous
origin that will be discussed later.
Epithelial lipids of interest to this study are free sphingoid bases because they
exhibit broad spectrum antimicrobial and antifungal activity (Bibel et al. 1992b, Bibel et
al. 1993, Bibel et al. 1995, Fischer et al. 2012, Fischer et al. 2013, Pavicic et al. 2007,
Payne et al. 1996, Possemiers et al. 2005). Concentrations of epithelial sphingoid bases,
sphingosines, dihydrosphingosines, and 6-hydroxysphingosines, range from 0.4 – 35.6
mg/g (Law et al. 1995b, Stewart and Downing 1995, Wertz et al. 1987, Wertz and
Downing 1989, Wertz and Downing 1990b), which is three orders of magnitude higher
than the levels of sphingoid bases measured in other human tissues (Law et al. 1995b).
Sphingosines and dihydrosphingosines present in stratum corneum are predominantly 18
carbons in length but range from 16 to 20 carbons (Wertz and Downing 1990b). Free
sphingoid bases are also present in saliva at concentrations ranging from 0.5 – 4.9 µg/ml
(Brasser et al. 2011a).
9
Sphingoid base structure and function
Sphingoid bases are long chain amino alcohols consisting of a hydrocarbon chain,
hydroxyl (-OH) groups, and an amine group. These amphipathic molecules vary in the
length of carbon chain, the degree of saturation, and the number of hydroxyl groups
present (Kendall and Nicolaou 2013). Four common sphingoid bases, sphingosine,
dihydrosphingosine, phytosphingosine, and 6-hydroxy-sphingosine form the basis of
different sphingolipids and can be released to exist as free sphingoid bases through the
action of hydrolytic enzymes (Law et al. 1995b).
Sphingosine, the most common sphingoid base in mammals, is present in oral
epithelium where a concentration gradient is evident, with higher sphingosine
concentrations in the stratum corneum and the superficial layer of non-keratinizing
regions (Law et al. 1995b). In addition to contributing to the permeability barrier of the
stratum corneum, sphingoid bases have many other cellular functions. Sphingosine
(Strum et al. 1997) and dihydrosphingosine (Darges et al. 1997) are strong inhibitors of
protein kinase C and therefore could contribute to communication between the stratum
corneum and the viable portion of the epidermis (Brogden et al. 2011). Sphingosine,
dihydrosphingosine, and phytosphingosine all affect inflammation (Darges et al. 1997,
Klee et al. 2007, Pavicic et al. 2007). Sphingosine derivatives such as sphingosine-1phosphate (Spiegel and Milstien 2003) and phytosphingosine-1-phosphate (Kim et al.
2007, Kim et al. 2006) act as signaling molecules.
The sphingoid bases of interest in this study are sphingosine, dihydrosphingosine,
and phytosphingosine because they exhibit potent antimicrobial activity against a broad
range of gram-positive and gram-negative bacterial species. Sphingosine (C18:1)
contains a single trans double bond between carbon-4 and carbon-5, hydroxyl groups on
carbon-1 and carbon-3, and an amino group on carbon-2. Dihydrosphingosine (C18:0) is
sphingosine’s saturated analog. Phytosphingosine (C18:0) is structurally similar to
dihydrosphingosine with the exception of a hydroxyl group at carbon-4.
10
Sebaceous lipids of the oral cavity
The most abundant lipids of the oral cavity are nonpolar lipids produced by
sebaceous follicles (Downing et al. 1993) which are found in all regions of the oral
mucosa and lining the vermillion border of the lips (Batsakis and el-Naggar 1990,
Downing et al. 1993, Gorsky et al. 1986, Olivier 2006). Sebaceous follicles are
functionally similar to pilosebaceous units but they lack terminal hairs that are associated
with major sebaceous glands of the skin. The outer basal cell layer of each sebaceous
follicle continuously proliferates, propelling immature sebocytes toward the center of the
gland. Lipids are synthesized in maturing sebocytes and each cell retains large amounts
of lipids. Sebocytes mature as they reach the center of the sebaceous gland where they
disintegrate, releasing their contents into the follicle (Downing et al. 1993, Smith and
Thiboutot 2008). Major lipid components of sebum include triglycerides, squalene, and
wax monoesters, along with lesser amounts of cholesterol and cholesterol esters (Brasser
et al. 2011b, Brogden et al. 2011, Downing et al. 1993). As sebum flows through the
follicular canal and onto the oral mucosal surfaces triglycerides are hydrolyzed to release
free fatty acids (Brogden et al. 2011, Drake et al. 2008, Thormar and Hilmarsson 2007).
Among the 12- to 18-carbon fatty acids derived from human sebum, sapienic acid
(C16:1∆6) is the major product and lauric acid (C12:0) is a minor product. Both sapienic
acid and lauric acid exhibit potent antimicrobial activity against a variety of grampositive and gram-negative bacteria (Bergsson et al. 2001a, Brogden et al. 2011, Fischer
et al. 2012, Fischer et al. 2013, Huang et al. 2011, Kabara et al. 1972b, Sado-Kamdem et
al. 2009). Sebaceous follicles are associated with the major salivary glands (Linhartova
1974, Martinez-Madrigal and Micheau 1989) and therefore salivary secretions can also
be a source of oral lipids (Brasser et al. 2011b). In fact, the total lipid fraction of saliva is
predominantly neutral lipids (95 – 99%) represented by free fatty acids, cholesterol,
cholesterol esters, and mono-, di-, and triglycerides (Defago et al. 2011, Larsson et al.
1996, Slomiany et al. 1985). Fatty acid concentrations in saliva range from 0.2 to 7.8
11
µg/ml (Brasser et al. 2011b, Palmerini et al. 2011). The recent discovery of wax esters
and squalene in saliva provides strong evidence that these lipids are of sebaceous origin
as these lipids are considered biochemical markers of human sebum (Brasser et al.
2011b).
Fatty acid structure and function
Fatty acids are also amphipathic molecules possessing a long hydrocarbon chain
with a terminal carboxylic acid group (-COOH). In biological systems fatty acids
typically have an even number of carbons, range from 10 to 28 carbons in length, and
differ in number and placement of double bonds. Fatty acids are important sources of
fuel, yielding large amounts of ATP when metabolized and are therefore crucial for
sustained contractile function of heart and skeletal muscle (Zhang et al. 2010). When not
bound to other compounds such as glycerol, sugars, or phosphate head groups, fatty acids
are termed “free fatty acids”. Sebaceous free fatty acids range from 7- to 22-carbons in
length but the major constituents of sebum are 12- to 18- carbons in length (Brogden et
al. 2011).
General trends in the antimicrobial activity of fatty acids reveal that this activity is
a function of carbon chain length as well as the presence, number, and orientation of
double bonds (Desbois and Smith 2010). Maximum antimicrobial activity of saturated
fatty acids is found in fatty acids 12-carbons in length (Brogden et al. 2011) but monounsaturated fatty acids of 14- to 16- carbons in length exhibit similar activity (Brogden et
al. 2011, Kabara et al. 1972b). Unsaturated fatty acids are generally more active than
saturated fatty acids of the same length (Kabara et al. 1972b, Zheng et al. 2005) and fatty
acids with cis double bonds are more active than those with trans double bonds (Galbraith
et al. 1971, Kabara et al. 1972b).
Of particular interest to this study are sapienic acid and lauric acid, which both
exhibit potent antimicrobial activity. Lauric acid is a straight-chain saturated 12-carbon
12
fatty acid. Sapienic acid (C16:1∆6) is a straight-chain fatty acid with a cis double bond at
carbon six which causes the molecule to twist, shrinking it to about the same physical
size as a 14-carbon fatty acid. Sebaceous sapienic acid also has a two-carbon extension
product (C18:1∆8) which does not exhibit any antimicrobial activity (Brogden et al.
2011).
Research Aims
Although sphingoid bases and fatty acids are present in oral mucosa and saliva
their antimicrobial activity on oral bacteria and effects on host cells have not been
explored. Therefore, the goal of this project was to determine the antimicrobial activity
of oral mucosal and salivary lipids against P. gingivalis, explore the mechanisms by
which oral lipids may inhibit or kill P. gingivalis, and examine their cytotoxic effects on
select host cells. To delineate the antimicrobial activity of oral lipids against P.
gingivalis, I developed experimental methodologies through the exploration of the
antimicrobial activity of sphingoid bases and fatty acids against a variety of grampositive and gram-negative bacteria that were aero-tolerant and easy to grow. Because
the antimicrobial lipids present in the oral cavity are also present on the skin, I chose to
study bacterial representatives endogenous to the skin and the oral cavity where they
would naturally be in contact with the lipids used for this study.
My first specific aim was to delineate the antimicrobial activity of oral
sphingoid bases and fatty acids against a variety of gram-positive and gramnegative bacteria endogenous to the oral cavity and skin (Chapter 2). For this study, I
established minimum inhibitory concentrations (MIC), minimum bactericidal
concentrations (MBC), and kill kinetics of each lipid for a variety of bacteria that are
present on the skin or in the oral cavity.
For my second specific aim: explore the morphological effects of oral lipids
against a representative gram-positive and gram-negative bacterium (Chapter 3), I
13
chose a gram-positive and a gram-negative bacterial representative from the first study to
explore the morphological effects of sphingoid base treatment through electron
microscopy. Because micrographs showed unique bacterial internal inclusion bodies in
both bacterial species tested, I measured the uptake of lipids by bacterial cells. Finally, in
an attempt to identify the site of treatment lipid activity, I investigated methods of
measuring the association of treatment lipids with bacterial cell lipids, which culminated
in data that were unclear as to furthering our understanding of lipid-lipid interactions in
this context.
Specific aim 3 was to explore and characterize the antimicrobial activity of
oral sphingoid bases and fatty acids against P. gingivalis (Chapter 4). For this study, I
utilized all the methods developed in the first two studies to test the antimicrobial effects
of fatty acids and sphingoid bases against P. gingivalis and explore potential mechanisms
of action through determination of MIC, MBC, kill kinetics, lipid-lipid interactions, and a
study of protein expression in lipid-treated and untreated P. gingivalis.
Chapter 5 addresses specific aim 4: determine the effects of oral sphingoid
bases against dendritic cells (DC) at cytotoxic and non-cytotoxic concentrations. For
this study I chose to examine sphingoid base effects on DCs because they are the primary
immune cells that would come into contact with the epithelium. Using a live/dead stain,
flow cytometry, and confocal microscopy I assessed the cytotoxic effects of lipids in the
range of antimicrobial activity against P. gingivalis that falls within normal physiologic
concentrations as well as within non-cytotoxic ranges.
Combined, these studies show that oral mucosal and salivary lipids exhibit dosedependent antimicrobial activity that is both lipid specific and bacteria specific and
induces a bacterial response that will be discussed throughout the remainder of this
dissertation. Importantly, there is a concentration range of lipids that is both antibacterial
to P. gingivalis and non-cytotoxic to DCs, suggesting a potential therapeutic or
prophylactic use of oral mucosal lipids.
14
CHAPTER 2
ANTIBACTERIAL ACTIVITY OF SPHINGOID BASES AND FATTY
ACIDS AGAINST GRAM-POSITIVE AND GRAM-NEGATIVE
BACTERIA
Common sphingoid bases and fatty acids are involved in the physical barrier,
permeability barrier, and immunologic barrier functions of skin and oral mucosa
(Cameron et al. 2007, Jungersted et al. 2008). Epithelial layers contain ceramides, free
fatty acids, and cholesterol; sebaceous lipids at the skin surface include a complex
mixture of triglycerides, fatty acids, wax esters, squalene, cholesterol and cholesterol
esters; and saliva contains these same sebaceous and epithelial lipids (Brasser et al.
2011b, Jungersted et al. 2008, Proksch et al. 2008). These sebaceous lipids contribute to
i) the transport of fat-soluble antioxidants to the skin and mucosal surfaces, ii) the proand anti-inflammatory properties of skin and mucosal surfaces, and iii) the innate
antimicrobial activity of the skin and mucosal surfaces (Smith and Thiboutot 2008,
Zouboulis 2004, Zouboulis et al. 2008).
Although the composition, biosynthesis, secretion, and function of cutaneous
lipids are well characterized from extensive and eloquent work done in the 1970s, little is
known about their role in controlling microbial infection and colonization. Certain fatty
acids and sphingoid bases found at the skin and mucosal surfaces are known to have
antibacterial activity and are thought to play a more direct role than previously realized in
innate immune defense against epidermal and mucosal bacterial infections (Drake et al.
2008). These include free sphingosines, dihydrosphingosines, lauric acid, and sapienic
acid. In human subjects, for example, the number of Staphylococcus aureus colony
forming units per unit area of skin is inversely proportional to both the sapienic acid
content and the free sphingosine content (Arikawa et al. 2002, Takigawa et al. 2005).
The lowest concentrations of both these antimicrobial lipids were found in subjects with
atopic dermatitis, for whom S. aureus infections are frequently a problem.
15
More recently, these same lipids have been shown to be present in the oral cavity,
in saliva, and at mucosal surfaces (Brasser et al. 2011a, Brasser et al. 2011b). The fatty
acids are derived from sebaceous triglycerides, while sphingoid bases are derived from
epithelial sphingolipids through the action of hydrolytic enzymes.
In this study, we hypothesized that the sphingoid bases sphingosine,
dihydrosphingosine, and phytosphingosine and the fatty acids sapienic acid and lauric
acid, commonly found on skin and mucosa, have antimicrobial activity against grampositive and gram-negative bacteria found on the skin and in the oral cavity. We also
suggest potential mechanisms for lipid antimicrobial activity and present their potential
as pharmaceuticals to improve therapies for treatment and control of a wide variety of
cutaneous and mucosal infections and inflammatory disorders.
Materials and Methods
Bacterial species and growth conditions
Bacteria commonly found on the skin and in oral microbiomes were used (Grice
and Segre 2011, Zarco et al. 2011). Escherichia coli and Serratia marcescens were also
included to obtain information about typical gram-negative bacterial susceptibility and
resistance. E. coli ATCC 12795, S. aureus ATCC 29213, S. marcescens ATCC 14756,
and Pseudomonas aeruginosa ATCC 47085 were grown for three hours in Mueller
Hinton broth (MHB; Difco Laboratories, Detroit, MI) at 37ºC. Corynebacterium bovis
ATCC 7715, Corynebacterium striatum ATCC 7094, and Corynebacterium jeikium
ATCC 43734 were grown for three hours in Brain Heart Infusion Broth (Difco
Laboratories, Detroit, MI) supplemented with 0.1% Tween 80 (ICN Biomedicals, Aurora,
Ohio) at 37ºC in an atmosphere containing 5% CO2. Streptococcus sanguinis ATCC
10556 and Streptococcus mitis ATCC 6249 were grown for three hours in tryptic soy
broth (TSB; Difco Laboratories, Detroit, MI) supplemented with 0.6% yeast extract
(Difco Laboratories, Detroit, MI) at 37ºC in an atmosphere containing 5% CO2. F.
16
nucleatum ATCC 25586 was grown in Schaedler’s- broth (Difco Laboratories, Detroit,
MI) for three hours at 37ºC in an anaerobic Coy Chamber (Coy Laboratory Products Inc.,
Grass Lake, MI). Before use, all bacterial cell suspensions were adjusted to contain 1 ×
108 CFU/ml (0.108 O.D., 600 nm, Spectronic 20D+, Thermo Fisher Scientific, Inc.,
Waltham, MA) and diluted with appropriate media to 107 CFU/ml (F. nucleatum), 106
CFU/ml (S. mitis), or 105 CFU/ml (remaining bacteria).
Preparation of lipids
D-sphingosine (e.g. sphingosine), phytosphingosine, D-erythrodihydrosphingosine (e.g. dihydrosphingosine), and lauric acid were obtained from Sigma
Chemical Company (St Louis MO). Sapienic acid was obtained from Matreya Inc.
(Pleasant Gap, PA). Lipids were dissolved in a chloroform:methanol solution (2:1), and
purity was confirmed by thin layer chromatography (TLC). Lipids, dried under nitrogen,
were then added to sterile 0.14 M NaCl to make a 1.0 mg/ml stock solution, and
sonicated for 30 minutes in a 37° C bath sonicator (Branson 2200, Hayward, CA) in five
minute increments to suspend the lipid. Lipids were then diluted to the desired
concentration using 0.14 M NaCl.
Antimicrobial assays
Broth microdilution assays were used to determine the MIC (minimum inhibitory
concentration; defined as the lowest concentration of lipid that reduced growth by more
than 50%) and the MBC (minimum bactericidal concentration; defined as the lowest
concentration of lipid that killed all bacteria) of each lipid for each bacterium (Kalfa et al.
2001, Turner et al. 1998). Briefly, lipid suspensions were diluted in 0.14 M NaCl (500 to
1 g/ml) in microtiter plates (Immunolon 1 microtiter plates, Thomas Scientific,
Swedesboro, NJ). Bacterial cultures in their respective concentrations and media were
then added. Media without microorganisms was added to 0.14 M NaCl in wells used as
the plate blank and negative control. Media with microorganisms was added to 0.14 M
17
NaCl in wells used as a positive growth control. After appropriate incubation times, the
optical density was read in the spectrophotometer (Spectromax Microplate Reader,
Molecular Devices Corp., Sunnyvale, CA) and the MIC was determined. At higher
concentrations, lipids had an optical density that interfered with the determination of an
MIC. Therefore, MBCs were also derived by plating bacteria from the completed broth
microdilution assays onto 5% sheep blood agar plates (Remel, Lenexa, KS) and
examining for the presence of colonies. MICs and MBCs were repeated in quadruplicate.
The sheep myeloid antimicrobial protein, SMAP28, (RGLRRLGRKIAHGVKK
YGPTVLRIIRIA-(NH2)) was synthesized as previously described (Kalfa et al. 2001) by
NeoMPS, Inc. (San Diego, CA) and suspended in 0.14 M NaCl. SMAP28 was included
in this study as a positive control to show that the microdilution assay was set up properly
and MICs were accurate and within previously reported ranges. SMAP28 is effective
against gram-positive bacteria, gram-negative bacteria, and fungi, but not against some
corynebacteria (Kalfa et al. 2001).
Killing kinetics assays
Killing kinetic assays were performed using the spiral plating method (Drake et
al. 1994). For this, a three-hour culture of each bacterial suspension, adjusted to the
appropriate concentration for each bacterium (described above), was split among five
groups and each was mixed with either 0.14 M NaCl (negative control) or lipids at a
contration equivalent to 10X MIC determined in the broth microdilution assays. At time
intervals of 0, 0.5, 1, 2, 3, 4, 6, 8, and 24 hours, one-ml samples of treated bacteria and
controls were removed, serially diluted into the appropriate media, and plated onto 5%
sheep blood agar plates (Remel, Lenexa, KS) using an Autoplate 4000 Automated Spiral
Plater (Advanced Instruments, Inc. Norwood, MA). Plates were incubated appropriately,
colonies were counted using standard spiral-plater methodology, and concentrations were
calculated. Killing kinetics assays were repeated in triplicate.
18
Statistical analyses
The exact Kruskal-Wallis test was employed to detect differences in the MIC and
MBC values utilizing a 5% level of statistical significance. This nonparametric analog to
ANOVA was used due to modest sample sizes and violations of the normality
assumptions for parametric procedures. Significance probabilities were for the test of the
null hypothesis that the distribution of outcome values is the same for all the treatment
groups designated. Post-hoc pair wise comparisons were not performed due to modest
sample sizes.
Two measures of killing kinetics were computed and analyzed. Trapezoidal area
under the curve (AUC) was used as a summary measure of bacterial viability over the
treatment time course, and comparisons were made with and without the inclusion of the
AUCs from the control sample. Significance probabilities reported are associated with
the null hypothesis that the distribution of trapezoidal area is the same among the
specified treatment groups. A second summary measure of killing kinetics over time
considered was time to zero, defined as the first time point at which total bacterial counts
reached zero. Note that, for certain of these longitudinal assays (ie. from a given vial),
none of the bacterial counts in the series reached zero. In such instances, the value of the
corresponding time to zero was assigned the highest rank for purposes of analysis. If
several such instances occurred in a given analyses, ties for the highest rank were
assigned.
Results
Sphingoid bases and fatty acids had antimicrobial activity for a variety of grampositive and gram-negative bacteria. MIC, MBC (Table 2.1), and kinetic killing curves
(Figure 2.1A-F) clearly showed that some sphingoid bases and fatty acids were more
potent for some microbial species than others. For example, sphingoid bases were
antimicrobial for two of the four gram-negative organisms tested: E. coli and F.
19
nucleatum (MIC range 0.7 to 15.6 µg/ml), while fatty acids were only active for F.
nucleatum (MIC range 2.1 to 6.5 µg/ml). Kinetic assays showed that killing of E. coli
and F. nucleatum with sphingosine and phytosphingosine occurred within 0.5 to 2 hours
(Table 2.2), whereas killing of F. nucleatum with lauric acid was more gradual and
occurred within 24 hours. Time to zero outcomes indicated significant differences
among lipid treatments for F. nucleatum (p = 0.0143). SMAP28 was used as a positive
assay control and MIC values ranged from 0.1 µg/ml for C. striatum and C. jeikeium to
10.0 µg/ml for S. marcescens.
Sphingoid bases were antimicrobial for all six of the gram-positive bacteria (MIC
range 0.3 to 13.0 µg/ml) (Table 2.1) and fatty acids were more active for oral
streptococcus species (MIC range 10.4 to 140.2 µg/ml) than S. aureus (MIC range 250 to
>500 µg/ml). Of the fatty acids, only lauric acid was weakly antibacterial for C. bovis, C.
striatum, and C. jeikeium (MIC range 208.3 to 416.7 µg/ml). Kinetic assays showed that
killing of S. aureus, S. sanguinis, S. mitis, and C. striatum with sphingosine and
phytosphingosine occurred within 0.5 to six hours (Table 2.2) but killing of S. aureus
with lauric acid and killing of S. sanguinis and S. mitis with sapienic acid was gradual
and occurred within 24 hours. Time to zero outcome comparisons indicated significant
differences among lipid treatments for S. mitis and C. striatum (p = 0.0036 for each).
Exact Kruskal-Wallis tests confirmed differences among the lipid treatments (p <
0.0001) for each of the bacterial species with the exception of S. marcescens and P.
aeruginosa (Table 2.1). Comparisons of the trapezoidal AUC also showed significant
differences among all treatment lipids for each of the organisms (p < 0.004 in all
instances) (Table 2.3). When controls were omitted from the analysis, significant
differences were seen among all the lipid treatments compared except for E. coli, where
there was no evidence that the AUC distribution differed for phytosphingosine and
sphingosine.
20
It is also worth noting that when bacteria were suspended in a simple saline
solution, kill kinetic assays were vastly different (data not shown). Complete killing of
E. coli and S. aureus, suspended in 0.14 M NaCl with phytosphingosine occurred within
0.5 hours. This was a reduction of 3 × 104 CFU/ml for E. coli and 2 × 104 CFU/ml for S.
aureus.
Discussion
Lipids typically found on the skin and mucosa have antimicrobial activity against
gram-positive bacteria and gram-negative bacteria found on the skin and in the oral
cavity. In this study, we show that the sphingoid bases sphingosine, phytosphingosine,
and dihydrosphingosine as well as two fatty acids, sapienic acid and lauric acid, had
variable antimicrobial activity for a variety of gram-positive and gram-negative bacteria.
These results are similar to that of others who have shown that sphingosine,
dihydrosphingosine, and phytosphingosine are active against Candida albicans (Bibel et
al. 1993) and fatty acids and their monoglycerides are antimicrobial for Group A and
Group B Streptococcus (Bergsson et al. 2001b, Drake et al. 2008, Nakatsuji et al. 2009,
Thormar and Hilmarsson 2007).
Although the exact mechanism of lipid antimicrobial activity is not fully
understood, there are a few possibilities to pursue. First, antimicrobial lipids may
penetrate and disrupt the cell wall layer of bacteria. In a recent study, we observed that
sphingoid bases appeared to lyse S. aureus, but not E. coli (Bratt et al. 2010a). After
incubation with sphingoid bases, preparations of S. aureus contained lysed cells and
identifiable fragments of the cell wall. Second, antimicrobial lipids may alter the
cytoplasmic membrane of bacteria. Bergsson et al. observed that fatty acids disrupted
and disintegrated the cytoplasmic membrane of C. albicans (Bergsson et al. 2001b). We
also observed that sphingoid bases appeared to alter the cytoplasmic membrane of S.
aureus, but not E. coli (Bratt et al. 2010a). Third, it is also possible that antimicrobial
21
lipids may directly penetrate the cell wall and cytoplasmic membrane of bacteria, enter,
and disrupt cytoplasmic contents similar to that described by Bergsson et al. for S. aureus
(Bergsson et al. 2001a).
The extent to which microorganisms can metabolize sphingoid bases and fatty
acids is not well known. It is possible that concentrations of lipids below the MIC can be
tolerated and metabolized and concentrations of lipids above the MIC cannot. It is also
possible that the bacteria used in this study can transport these lipids into the cell,
accumulating as intracellular inclusions. We recently observed that sphingoid bases
induced the formation of intra-cytoplasmic inclusions (Bratt et al. 2010a). Whether these
inclusions are composed of accumulated lipids or bacterial-derived proteins is not yet
known and is under investigation.
The high antimicrobial activity of lipids suggests that they may have applications
as therapies to prevent or treat a wide variety of skin infections. These lipids are easy to
obtain, have potent antimicrobial activities, and are likely to have low toxicity. In
addition to direct antibacterial action, antimicrobial peptides are also chemotactic and can
attract leukocytes to sites of infection (Dale 2002, Gallo et al. 2002). The sphingoid
bases are also inhibitors of protein kinase C and can thereby modulate many biochemical
actions. In addition, free sphingosine can be phosphorylated to produce sphingosine-1phosphate which is a potent bioactive metabolite that regulates diverse processes of
importance to inflammation and immunity (Spiegel and Milstien 2011).
Phytosphingosine may be an ideal candidate for treating acne vulgaris (Klee et al.
2007, Pavicic et al. 2007) as it has been shown to be antimicrobial for Propionibacterium
acnes in vitro; down-regulates the pro-inflammatory chemokines IL-8, CXCL2, and
endothelin-1 in primary human keratinocytes; reduces the release of both lactate
dehydrogenase and IL-1 in response to sodium dodecyl sulfate; is anti-inflammatory
when tested in an organotypic skin model; and enhances the resolution of acne when
applied topically. Lauric acid (C12:0) has promise as a potential therapeutic for the
22
treatment of acne because it has MICs over 15 times lower than those of benzoyl
peroxide and is not cytotoxic in vitro to human sebocytes or in vivo in mouse dermis
(Nakatsuji et al. 2009).
Lipids common to the skin and oral cavity, sphingosine, phytosphingosine,
dihydrosphingosine, sapienic acid, and lauric acid, had variable antimicrobial activity for
a variety of gram-positive and gram-negative bacteria. Fatty acids and sphingoid bases
may be contributing to defensive barrier functions of the skin and oral cavity and may
have potential for prophylactic or therapeutic intervention of infection.
Sphingosine
Phytosphingosine
Dihydrosphingosine
Lauric acid
Sapienic acid
E. coli
MIC mean
MBC mean
MBC median
7.8 ± 0.0
19.6 ± 13.6
19.6
3.9 ± 0.0
15.6 ± 0.0
15.6
5.6 ± 0.0
39.1 ± 15.6
31.3
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
P. aeruginosa
MIC mean
MBC mean
MBC median
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
p-value
SMAP28c
p<0.0001b
1.7 ± 0.7
ND
ND
1.6 ± 0.6
ND
ND
Table 2.1. Inhibitory and bactericidal activity of sphingoid bases and fatty acids for gram-negative and gram-positive bacteria.
a
MIC or MBC values are larger than the upper limit of detection for the assay.
b
c
Denotes significance at the 0.05 level. Significance probabilities associated with the nonparametric Kruskal-Wallis test of the null
hypothesis that the distribution of MBC values is the same across all treatment groups with a specified bacterial species.
SMAP28 was used as a positive assay control to show that the microdilution assays were set up properly and MICs were accurate and
within previously reported ranges. SMAP28 MBCs were not completed and results were not included in statistical analysis.
Note: Gram-negative bacteria include: E. coli, P. aeruginosa, S. marcescens, and F. nucleatum. Gram-positive bacteria include: S.
aureus, S. sanguinis, S. mitis, C. bovis, C. striatum, C. jeikeium. Data show mean MIC and MBC (µg/mL) ± standard deviation.
ND = not done.
23
Sphingosine
Phytosphingosine
Dihydrosphingosine
Lauric acid
Sapienic acid
S. marcescens
MIC mean
MBC mean
MBC median
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
3.3 ± 1.4
ND
ND
F. nucleatum
MIC mean
MBC mean
MBC median
0.7 ± 0.2
4.9 ± 2.0
3.9
3.3 ± 0.7
3.9 ± 0.0
3.9
2.0 ± 0.2
2.0 ± 0.0
2.0
2.1 ± 1.0
6.8 ± 2.0
7.8
6.5 ± 1.3
86.0 ± 46.9
93.8
p<0.0001b
0.6 ± 0.1
ND
ND
S. aureus
MIC mean
MBC mean
MBC median
1.3 ± 0.3
1.3 ± 0.5
1.0
1.6 ± 0.3
7.8 ± 0.0
7.8
1.3 ± 0.3
4.7 ± 3.7
4.9
250.0 ± 0.0
250.0 ± 0.0
250.0
>500.0 ± 0.0a
62.5 ± 0.0
62.5
p<0.0001b
3.3 ± 1.3
ND
ND
S. sanguinis
MIC mean
MBC mean
MBC median
2.0 ± 0.0
1.3 ± 0.5
1.0
7.8 ± 0.0
3.4 ± 1.0
3.9
0.7 ± 0.0
1.3 ± 0.5
1.0
10.4 ± 2.6
125.0 ± 0.0
125.0
52.1 ± 10.4
31.3 ± 0.0
31.3
p<0.0001b
5.0 ± 0.0
ND
ND
S. mitis
MIC mean
MBC mean
MBC median
0.5 ± 0.2
0.2 ± 0.0
0.2
7.8 ± 3.9
3.0 ± 1.1
3.0
0.3 ± 0.0
0.3 ± 0.2
0.2
15.6 ± 0.0
15.6 ± 11.1
11.7
140.2 ± 20.8
375.0 ± 144.3
375.0
p<0.0001b
5.0 ± 0.0
ND
ND
SMAP28c
24
Table 2.1. Continued
p-value
Sphingosine
Phytosphingosine
Dihydrosphingosine
Lauric acid
Sapienic acid
C. bovis
MIC mean
MBC mean
MBC median
1.6 ± 0.3
15.6 ± 0.0
15.6
5.2 ± 1.3
62.5 ± 0.0
62.5
5.2 ± 1.3
15.6 ± 0.0
15.6
416.7 ± 83.3
156.3 ± 62.5
125.0
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
C. striatum
MIC mean
MBC mean
MBC median
1.3 ± 0.3
2.0 ± 0.0
2.0
4.2 ± 1.3
7.8 ± 0.0
7.8
1.0 ± 0.0
2.0 ± 0.0
2.0
250.0 ± 0.0
375.0 ± 144.3
375.0
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
C. jeikeium
MIC mean
MBC mean
MBC median
5.2 ± 1.3
11.7 ± 4.5
11.7
13.0 ± 5.2
31.3 ± 0.0
31.3
10.4 ± 2.6
15.6 ± 0.0
2.0
208.3 ± 41.7
93.8 ± 36.1
93.8
>500.0 ± 0.0a
>500.0 ± 0.0a
NDa
p-value
SMAP28c
p<0.0001b
0.5 ± 0.2
ND
ND
p<0.0001b
0.02 ± 0.0
ND
ND
p<0.0001b
0.03 ± 0.0
ND
ND
Table 2.1. Continued
25
Bacterium
Treatment lipids compared
Median Time to Zero
(hours)
P-values
E. coli
Phytosphingosine
Sphingosine
2.0
0.5
0.10
F. nucleatum
Phytosphingosine
Sphingosine
Sapienic Acid
1.0
0.5
8.0
0.0143a
S. aureus
Phytosphingosine
Sphingosine
24.0
1.0
0.10
S. sanguinis
Phytosphingosine
Sphingosine
Sapienic Acid
0.5
0.5
0.5
1.00
S. mitis
Phytosphingosine
Sphingosine
Lauric Acid
24.0
6.0
1.0
0.0036a
C. striatum
Phytosphingosine
Sphingosine
Lauric Acid
3.0
4.0
0.5
0.0036a
Table 2.2. Time to zero comparisons of lipid kill kinetics for gram-positive and gram negative bacteria.
a
Denotes significance at the 0.05 level.
26
Note: Comparisons were made without the control group as the control samples did not produce zero values. Significance
probabilities are associated with the exact non-parametric Kruskal-Wallis test of the null hypothesis that the distribution of
trapezoidal area is the same for all treatment groups designated.
Bacterium
AUC of bacteria
alone
Treatment lipids
compared
AUC of
treatment lipid
P-values
(including controls)
P-values
(excluding controls)
E. coli
249.08
Phytosphingosine
Sphingosine
3.82
0.75
0.0036a
0.10
F. nucleatum
108.13
Phytosphingosine
Sphingosine
Sapienic Acid
1.67
1.00
16.74
0.000065a
0.0036a
S. aureus
201.04
Phytosphingosine
Sphingosine
Lauric Acid
27.17
1.53
169.59
0.000065a
0.0036a
S. sanguinis
180.98
Phytosphingosine
Sphingosine
Sapienic Acid
0.63
0.45
0.40
0.00052a
0.0214a
S. mitis
210.11
Phytosphingosine
Sphingosine
Lauric Acid
2.09
14.79
46.43
0.000065a
0.0036a
Table 2.3. Trapezoidal AUC comparisons of lipid treatments as a summary measure of bacterial viability over the treatment
time course.
a
Denotes significance at the 0.05 level.
Note: Significance probabilities are associated with the exact nonparametric Kruskal-Wallis test of the null hypothesis that the
distribution of trapezoidal area is the same for all treatment groups designated.
27
Bacterium
AUC of bacteria
alone
Treatment lipids
compared
AUC of
treatment lipid
P-values
(including controls)
P-values
(excluding controls)
C. striatum
13.59
Phytosphingosine
Sphingosine
Lauric Acid
5.92
8.07
0.78
0.000065a
0.0036a
Table 2.3. Continued
28
29
Figure 2.1. Kill kinetics of lipid treatments against gram-positive and gram-negative
bacteria. Lipid treatments were all equal to 10X their MIC. Where no
bacteria were recovered, +1 was added to zero values before log
transformation of the data. Geometric mean of n=3 is shown for each data
point. Error bars show standard error of the mean (SEM). Gap in C. striatum
growth control (F) indicates no available information at those data points.
30
Figure 2.1. Continued
31
Figure 2.1. Continued
32
CHAPTER 3
SPHINGOID BASES INDUCE ULTRASTRUCTURAL DAMAGE IN
ESCHERICHIA COLI AND STAPHYLOCOCCUS AUREUS AND
ALTER THEIR LIPID COMPOSITION
Sphingoid bases and neutral lipids are present in the stratum corneum where they
likely contribute to the permeability and innate immunologic barriers of the skin and oral
mucosa (Brogden et al. 2012, Jungersted et al. 2008). Included among these lipids are
fatty acids, derived from sebaceous triglycerides, and free sphingosine and
dihydrosphingosine, derived from epithelial sphingolipids via hydrolytic enzymes.
Although the antibacterial activity of these common lipids against both gram-positive and
gram-negative bacteria has been established the mechanisms of action have not yet been
established.
Sphingosine, phytosphingosine, and dihydrosphingosine are all similar in
structure. Sphingosine (C18:1) is a long chain unsaturated fatty alcohol with a single
trans double bond between C4 and C5, hydroxyl groups on C1 and C3, and an amino
group on C2. Dihydrosphingosine (C18:0) is sphingosine’s saturated analog. Both
mediate a variety of cellular processes (Bu et al. 2006, Saba and Hla 2004, Shi et al.
2007, Spiegel and Milstien 2003, Spiegel and Milstien 2011), through the inhibition of
protein kinase C (PKC) (Darges et al. 1997). Phytosphingosine (C18:0) is structurally
similar to dihydrosphingosine with the exception of a hydroxyl group at C4.
Phytosphingosine also mediates a variety of cellular processes and has anti-proliferative
and anti-inflammatory properties (Kim et al. 2006, Pavicic et al. 2007).
Sphingosine, dihydrosphingosine, and phytosphingosine exhibit varying degrees
of antimicrobial activity (Bibel et al. 1992b, Bibel et al. 1993, Drake et al. 2008, Klee et
al. 2007, Pavicic et al. 2007) and are both lipid-specific and microorganism-specific
against a variety of gram-positive and gram-negative bacteria (Bibel et al. 1992a, Bibel et
al. 1992b, Bibel et al. 1993, Fischer et al. 2012). Bibel and colleagues showed that these
33
sphingoid bases are highly active against gram-positive bacteria and fungi, but relatively
inactive against gram-negative bacteria. Based on studies including L-forms of S.
aureus, the site of activity was suggested to be the cell wall. Electron microscopy
showed cell wall lesions, disruption of the membrane, and leakage of cellular content
(Bibel et al. 1993).
Recently, we found that sphingosine, dihydrosphingosine, and phytosphingosine
are active (MIC range 0.7 – 31.3 µg/ml) against E. coli, S. aureus, C. bovis, C. striatum,
C. jeikium, S. sanguinis, S. mitis, and F. nucleatum but not against S. marcescens and P.
aeruginosa (MIC >500 µg/ml) (Fischer et al. 2012). Kinetics assays revealed that
complete killing is achieved in as little as 0.5 h for some lipid-bacteria combinations but
up to 24 h are required for other combinations. Although the antibacterial activity of
these common sphingoid bases against both gram-positive and gram-negative bacteria
has been established, the mechanisms of action have not yet been established.
In this study, we begin to assess lipid activity against a representative grampositive and gram-negative bacteria: S. aureus, an opportunistic skin pathogen
contributing to a wide variety of diseases leading to an estimated 478,000 hospitalizations
and 11,000 deaths in the United States annually (Klein et al. 2007), and E. coli, another
contributor to skin and soft tissue infections (Dryden 2010, Petkovsek et al. 2009).
Similar to the results of Bibel and colleagues, we show that sphingoid bases induce
ultrastructural damage. Furthermore, we show that 1) sphingoid bases accumulate in the
bacterial cell; 2) sphingoid bases induce differential ultrastructural changes in
representative gram-positive and gram-negative bacteria; and 3) sphingoid bases induce
the presence of intracellular inclusions. The combination of these ultrastructural changes
indicates a need for further study into potential mechanisms for their antimicrobial
activity against microorganisms.
34
Materials and Methods
Bacterial species and growth conditions
E. coli ATCC® 12795TM and S. aureus ATCC® 29213TM were grown for three
hours in MHB (Difco Laboratories, Detroit, MI) at 37ºC. Bacterial cell suspensions were
adjusted to contain 1 × 108 CFU/ml (0.108 O.D., 600 nm, Spectronic 20D+, Thermo
Fisher Scientific, Inc., Waltham, MA). For scanning electron microscopy (SEM),
suspensions were serially diluted to 1 × 105 CFU/ml before treatment and for
transmission electron microscopy (TEM), suspensions remained at 1 × 108 CFU/ml.
Preparation of lipids
Sphingosine, dihydrosphingosine, and phytosphingosine were obtained from
Sigma Chemical Company (St Louis MO) and prepared as described in Chapter 2.
Prepared stock solutions of 1.0 mg/ml were diluted to the desired concentrations using
0.14 M NaCl.
Preparation of lipid-damaged bacterial cells
Broth cultures of E. coli and S. aureus were incubated with sphingosine,
dihydrosphingosine, or phytosphingosine at 10X the previously determined MIC (Fischer
et al. 2012) for 0.5 hours (E. coli treatments) and four hours (S. aureus treatments) at
37°C. E. coli was treated with 39 µg/ml phytosphingosine, 104 µg/ml sphingosine, or
312 µg/ml dihydrosphingosine. S. aureus was treated with 13 µg/ml phytosphingosine,
16 µg/ml sphingosine, or 20 µg/ml dihydrosphingosine. In order to visualize cells in
various stages of death, incubation times were based on killing kinetics (Fischer et al.
2012) so that each suspension contained both viable (<50%) and non-viable (>50%) cells.
Scanning electron microscopy
After treatment with lipids, E. coli and S. aureus were layered on a nucleopore
membrane (SPI Supplies, West Chester, PA), fixed with 2.5% glutaraldehyde in 0.1 M
35
sodium cacodylate buffer, pH 7.4, for one hour in an ice bath, and washed twice in 0.1 M
sodium cacodylate buffer, pH 7.4, for four minutes. Samples were then further fixed in
1% osmium tetroxide for 30 minutes, washed twice in double distilled water, and
dehydrated in a series of 25%, 50%, 75%, 95%, and absolute ethanol solutions followed
by hexamethyldisilizane. Membranes containing E. coli or S. aureus were then mounted
on stubs, sputter coated with gold and palladium, and examined with a Hitachi S-4800
SEM (Hitachi High-Technologies Canada, Inc., Toronto, Ontario Canada).
Transmission electron microscopy
After treatment with lipids, E. coli and S. aureus were fixed in 2.5%
glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for one hour in an ice bath,
and washed twice in 0.1 M sodium cacodylate buffer, pH 7.4, for 20 minutes. After
fixation, the organisms were pelleted by centrifugation and suspended in warm 0.9%
agarose in 0.1 M sodium cacodylate buffer, pH 7.4. The agarose was diced into one-mm
cubes and washed twice in 0.1 M sodium cacodylate buffer, pH 7.4, for 20 minutes.
Cubes of agarose containing treated E. coli or S. aureus were then treated with 1%
osmium tetroxide for one hour, washed twice in 0.1 M sodium cacodylate buffer, pH 7.4,
for 20 minutes, dehydrated in a series of 30%, 50%, 70%, 95%, and absolute ethanol
solutions, cleared in propylene oxide, infiltrated in a propylene oxide-Epon mixture (1:1),
embedded in Epon, and polymerized at 60oC for 48 hours. Ultrathin sections were cut,
placed on formvar-coated nickel grids, and then stained with 5% uranyl acetate and
Reynold’s lead citrate. Samples were examined in a JEOL JEM-1230 TEM (JEOL USA,
Inc., Peabody, MA USA).
Cell dimensions and statistical analysis
Cells were randomly selected and photographed. To quantitate the observed
effects of lipid treatments on cells, we measured cell dimensions using ImageJ. For E.
coli, length (L) and width (W) measurements were taken across the approximate center of
36
each bacterium. For S. aureus, vertical diameter measurements (d) were taken across the
approximate center of each cell. Analyses are based on a minimum of 10 measurements
for each treatment/organism combination.
Visible surface areas of E. coli (L × W) and S. aureus (π r2, where r = d/2) were
computed as a method of examining treatment-induced change in overall bacterial size.
The nonparametric Kruskal-Wallis test was employed with a 5% level of significance to
test the null hypothesis that the distribution of visual surface areas is the same for all the
treatment groups designated. Pairwise comparisons among the four treatment groups
within each bacterial species were performed using the exact Wilcoxon Rank Sum test.
The Bonferroni correction was used to adjust for multiple comparisons to maintain an
overall Type I Error level of 5%.
Isolation of lipids from bacteria
Broth cultures of E. coli and S. aureus were incubated with each of 0.14 M NaCl
(negative control), sphingosine, dihydrosphingosine, and phytosphingosine at 500 µg/ml
(total volume of treatment was 5 ml per sample) for four hours at 37°C. Sodium azide
(0.05%) was added to kill the bacteria before pelleting by centrifugation. Whole cell
pellets were frozen at -80°C, lyophilized, and lipids extracted using a previously
described method (Wertz et al. 1987) that consisted of successive extractions with
chloroform:methanol mixtures (2:1, 1:1, and 1:2) for two hours each at room
temperature. Extracted lipids were recovered by evaporation of the solvent under a
stream of nitrogen. The lipids were then redissolved in five ml chloroform:methanol
(2:1) and washed with one ml 2 M potassium chloride to remove salts and other water
soluble materials (Folch et al. 1957). The resulting upper phase was discarded and the
lower phase was again dried under nitrogen. Dried lipids were weighed and reconstituted
in chloroform:methanol, 2:1 at a concentration of 10 mg/ml. Additional controls
included suspensions of each treatment lipid in bacterial growth media (MHB) without
37
bacteria followed by centrifugation and resuspension in chloroform:methanol (2:1) to test
the ability of the sphingoid bases to sediment.
Lipid analysis
The lipids from each treatment and control were analyzed for sphingoid bases by
quantitative TLC (QTLC) as previously described (Wertz and Downing 1989).
Chromatograms were developed with chloroform:methanol:water (40:10:1). Developed
chromatograms were sprayed with 50% sulfuric acid and charred by heating slowly to
220°C on a hotplate. Digital images were obtained using a Hewlett-Packard Scanjet
3500c and analyzed using TNIMAGE (Thomas Nelson, Bethesda, MD) in strip
densiometry mode to estimate the total extracted lipid weight in each of the treated and
untreated bacterial samples as well as controls. Percentage of lipid uptake for each
sample was calculated by dividing the total extracted lipid weight by the total weight of
lipid added. Sphingosine served as a standard for quantification (Weerheim and Ponec
2001).
Results
Scanning electron microscopy
Control E. coli were 2.04 ± 0.46 µm long (mean ± standard error of the mean) by
0.63 ± 0.03 µm wide in size and exhibited typical rod morphology (Henk et al. 1995). In
contrast, E. coli treated with sphingoid bases were distorted and their surfaces were
concave and rugate (Figure 3.1). Many cells had external blebs. E. coli treated with
sphingoid bases were notably smaller in length and width when compared with the
controls. E. coli treated with sphingosine were 1.33 ± 0.24 µm long by 0.52 ± 0.03 µm
wide; dihydrosphingosine were 1.23 ± 0.29 µm long by 0.59 ± 0.07 µm wide; and
phytosphingosine were 1.82 ± 0.73 µm long by 0.73 ± 0.03 µm wide in size (Table 3.1).
38
The Kruskal-Wallis test for differences in the visible surface areas of untreated
and sphingoid base-treated E. coli (p<0.0001) was significant at the 5% level of
significance, indicating that the distribution of visual surface areas differed among the
treatment groups. Post-hoc pairwise comparisons of treatment for E. coli and the
associated treatments (Table 3.2) indicated that dihydrosphingosine and sphingosine had
distributions of visual surface areas that significantly differed from the control
(p=<0.0001 for both) after multiple comparisons adjustments using an overall 5% level of
significance. They did not, however, significantly differ from each other (p=0.6784).
There was no evidence that the distribution of visual surface areas differed between
phytosphingosine and the control. Dihydrosphingosine and sphingosine were each
significantly different from phytosphingosine (p <0.0001 for both) after multiple
comparisons adjustment. Both had medians which are less than that observed with
phytosphingosine treatment.
Control S. aureus were 0.74 ± 0.03 µm in diameter (± SEM) and exhibited typical
gram-positive coccus morphology and staphylococcal arrangement. S. aureus treated
with sphingoid bases were also distorted to various degrees. Some cells were concave
and rugate and appeared to be in various stages of lysis with compromise of the cell wall
and plasma membrane. Some cells in clusters were lysed leaving remnants of the cell
wall and cellular debris near adjacent cells (Figure 3.1). Septal grooves appeared to be
more pronounced and deeper than in the control S. aureus cells. S. aureus treated with
sphingosine were 0.59 ± 0.08 µm in diameter; cells treated with dihydrosphingosine were
0.57 ± 0.07 µm in diameter; while cells treated with phytosphingosine were 0.73 ± 0.12
µm in diameter (Table 3.1).
The Kruskal-Wallis test for S. aureus (p<0.0001) was significant at the 5% level
of significance, indicating treatment differences in the distribution of visual surface area.
After adjustment for multiple comparisons using the Bonferroni method, post-hoc
pairwise comparisons among S. aureus and the related treatments (Table 3.3) also
39
indicated that dihydrosphingosine and sphingosine had smaller visible surface areas that
were different from the control (p<0.0001 for both treatments). They did not, however,
significantly differ from each other (p=0.2993). There was no evidence that the
distribution of visual surface areas differed between phytosphingosine and control.
Additionally, dihydrosphingosine and sphingosine were each significantly different from
phytosphingosine (p <0.0001 for both). Both had medians which were less than that of
phytosphingosine.
Transmission electron microscopy
In thin sections, control E. coli exhibited typical gram-negative rod morphology
(Bayer and Thurow 1977, Bayer et al. 1985) and the outer envelope, interspace, and
cytoplasmic membrane were visible (Figure 3.2). In E. coli treated with sphingoid bases,
the outer envelope and cytoplasmic membrane appeared intact and there was no visual
evidence that the bacterial cell walls or membranes were damaged.
Interestingly, in E. coli treated with sphingoid bases, there were obvious electron
dense intracellular inclusion bodies of various sizes and shapes. In many instances, these
bodies filled the intracellular content of the cells. The remaining cytoplasm was not
uniform suggesting aggregation or flocculation of intracellular contents.
In thin sections, S. aureus had a typical gram-positive morphology (Harder et al.
2001, Shimoda et al. 1995) and the cell wall and cytoplasmic membrane were clearly
visible. The cytoplasm had a characteristic uniform granularity with an occasional
fibrinous-like whirl characteristic of the nucleoid (Figure 3.2).
S. aureus cells treated with sphingoid bases were in various stages of
disintegration and lysis (Figure 3.2). Some cells had intact cell walls and cytoplasmic
membranes but the cells contained a flocculated cytoplasm. Septal grooves appeared to
be more pronounced and deeper than in the control S. aureus cells. Other cells had lysed
and there were cross sections of cell wall and cellular debris visible near damaged cells.
40
Similar to E. coli, S. aureus treated with sphingoid bases also contained electron
dense intracellular inclusion bodies of various sizes and shapes that filled the intracellular
content of the cells. In some cells there were additional vesicles. Whether these were
remnants of cytoplasmic membrane still within the cell wall shell are yet to be
determined. The remaining cytoplasm was not uniform also suggesting an aggregation or
flocculation of intracellular contents.
Thin layer chromatography
Both E. coli and S. aureus took up large amounts of sphingosine,
phytosphingosine, and dihydrosphingosine relative to controls (Figure 3.3). A small
amount of each of the sphingoid bases did remain on the test tube walls or sediment from
the medium upon centrifugation and was present in the control sphingosine,
phytosphingosine, and dihydrosphingosine lanes but this was a relatively small amount
compared to the lipids extracted from treated bacterial samples. Bacterial counts were
completed at the beginning and end of the treatment period and confirmed killing of both
E. coli and S. aureus by all three treatments. After the four-hour treatment period, E. coli
dropped from 3.6 × 108 CFU/ml to 5.2 × 102 upon treatment with sphingosine, 1.0 × 106
with phytosphingosine treatment, and 6.0 × 102 with dihydrosphingosine treatment. S.
aureus went from 2.2 × 108 CFU/ml in the untreated control to 5.0× 102 with sphingosine
treatment, 3.0 × 104 with phytosphingosine treatment, and 2.0 × 105 upon treatment with
dihydrosphingosine.
Discussion
An extensive number of host innate immune factors induce extensive
ultrastructural damage to gram-negative and gram-positive bacterial cells. These factors
include anionic peptides (Brogden et al. 1996), cathelicidins (Kalfa et al. 2001), and
defensins (Harder et al. 2001, Shimoda et al. 1995). We report here that sphingoid bases,
which have been shown to be antimicrobial by our group and others (Bibel et al. 1993,
41
Bibel et al. 1995, Payne et al. 1996), also induce extensive ultrastructural damage to E.
coli and S. aureus.
Treated cells from both species were distorted and, in some instances, were
notably smaller in size. Their outer surfaces were concave and rugate in appearance,
demonstrating damage to the cell. Treated cells of S. aureus also had a noticeable loss of
the cell wall. In thin sections of E. coli, there was no evident compromise of the bacterial
cell walls and membranes appeared to be intact. In S. aureus, there was obvious
disruption and loss of cell wall and membrane. Treated cells of both E. coli and S. aureus
contained unique internal inclusion bodies. Hence, lipids at the skin surface induce both
extracellular and intracellular damage.
Our results are similar to other studies of sphingoid bases against S. aureus in that
sphingosine and dihydrosphingosine interfere with cell wall synthesis (Bibel et al. 1993).
Dihydrosphingosine-treated S. aureus has multiple lesions in the cell wall, evaginations
in the plasma membrane, and a loss of ribosomes in the cytoplasm (Bibel et al. 1993).
The cell wall lesions may be sequelae of the affected plasma membrane. However, while
treatment of E. coli with sphingosine results in surface bleb formation, the cell wall
appears to be intact. Phytosphingosine appears to cause more overt cell wall damage.
Our results are also similar to those obtained when cells are treated with fatty
acids or monoglycerides, which do not disrupt the integrity of the bacterial cell. Often
there are no visible effects on bacterial cell walls by either SEM or in thin sections
examined by TEM. Rather, the site of action appears to be the plasma membrane, which
is often partially dissolved or missing. For example, Group B Streptococcus treated with
10 mM monocaprin for 30 minutes are killed by disintegration of the cell membrane,
leaving the bacterial cell wall intact (Bergsson et al. 2001a). The plasma membrane and
electron transparent granules are gone. Interestingly, there are no visible effects of
monocaprin on the bacterial cell wall directly. No changes can be seen by either SEM or
in thin sections examined by TEM. Similarly, C. albicans treated with capric acid
42
(C10:0) has a disrupted or disintegrated plasma membrane with a disorganized and
shrunken cytoplasm (Bergsson et al. 2001b). Again, no visible changes are seen in either
the shape or the size of the cell wall. Whether there is a general fluidizing effect resulting
in leakage of cellular contents or a more specific interaction with membrane components
is not yet known.
The exact mechanism of sphingoid base action on the bacterial cell is currently
being elucidated. All microorganisms have polar lipids in their cytoplasmic membranes
(gram-positive and gram-negative bacteria) and the inner leaflet of the outer membrane
(gram-negative bacteria) (Brogden 2009). It appears likely that lipids may insert into the
outer envelope and cytoplasmic membranes of gram-negative bacteria and the
cytoplasmic membranes of gram-positive bacteria. Direct changes in the physical
properties of the bacterial membranes resulting from sphingoid base insertion may render
the membrane non-functional and thus may be the basis for bactericidal activity.
Alternatively, lipids may penetrate and accumulate in the cytoplasm. We have
shown here that these sphingoid bases are taken up by both E. coli and S. aureus in large
quantities and there is a possibility that they may be contributing to the internal inclusions
seen in our micrographs. Microorganisms are known to accumulate lipid inclusions and
microcompartments of varying shapes and compositions. Triacylglycerol inclusions and
neutral storage lipid inclusions are two examples (Alvarez and Steinbuchel 2002,
Kalscheuer et al. 2007). In the case of sphingoid bases, the presence of these lipids may
interfere with cell metabolism. It is possible that the sphingoid bases may specifically
inhibit certain enzymes in a manner similar to that by which they inhibit mammalian
protein kinase C (Hannun et al. 1986).
It is interesting to note that sphingosine, dihydrosphingosine, and
phytosphingosine induced differing effects on E. coli and S. aureus. Pairwise
comparisons across lipid treatments and controls for each bacterium were significant
between the controls and each of dihydrosphingosine and sphingosine but not for
43
phytosphingosine. When comparing lipid treatments, phytosphingosine was different
from dihydrosphingosine and sphingosine, but sphingosine and dihydrosphingosine
showed no significant differences from each other. Sphingosine and dihydrosphingosine
have similar structures, differing by only a single trans double bond. The molecular
structure of phytosphingosine, however, contains an additional hydroxyl group, making it
more polar. This could also explain the high variability seen in the visual surface area
differences of both E. coli and S. aureus when treated with phytosphingosine. The
increased hydrophilicity of phytosphingosine may contribute to slower partitioning into
the bacterial membrane.
Increase in S. aureus skin colonization is associated with lipid deficiencies. For
example, both deficient hexadecanoic acid production (Takigawa et al. 2005) and
decreased levels of sphingosine (Arikawa et al. 2002) are associated with atopic
dermatitis and a subsequent increase in S. aureus skin colonization. Additionally, failure
to clear S. aureus skin infections within innate immunodeficient mice is linked to
mutation of an enzyme necessary for palmitic and oleic acid production (Georgel et al.
2005). Understanding the specific activities of lipids on bacteria contributes to
knowledge of the roles of lipids in the control of bacteria in the oral mucosa and on the
skin.
In this study, we show that sphingoid bases induce unique ultrastructural damage.
Sphingoid base-treated E. coli exhibited intact membranes and multiple internal inclusion
bodies. Sphingoid base-treated S. aureus had obvious membrane and cell wall damage as
well as multiple internal inclusion bodies. In conclusion, sphingoid bases commonly
found on skin and in mucosal secretions have differential antimicrobial activity against
gram-positive and gram-negative bacteria and may have potential for prophylactic or
therapeutic intervention of infection.
E. coli
(µm2)a
Treatments
S. aureus
(µm2) a
Nb
Mean (SD)
Medianc
Nb
Mean (SD)
Medianc
Dihydrosphingosine
13
0.663 (0.147)
0.658
25
0.218 (0.017)
0.128
Phytosphingosine
10
1.300 (0.291)
1.356
33
0.128 (0.033)
0.215
Sphingosine
11
0.681 (0.105)
0.684
30
0.215 (0.069)
0.135
Control
11
1.302 (0.318)
1.215
10
0.140 (0.038)
0.219
Table 3.1. Visual surface area descriptive statistics for untreated and sphingoid base-treated E. coli and S. aureus.
a
Visual surface areas of E. coli (L × W) and S. aureus (π r2, where r = d/2) were computed using measurements across the
approximate center of each bacterium as a method of examining treatment-induced change in overall bacterial size.
b
c
N = number of bacteria measured.
Kruskal-Wallis tests for differences in the visible surface areas of treated and sphingoid base-treated bacteria were significant
(p<0.0001) for E. coli and S. aureus but post-hoc pairwise comparisons varied (see tables 3.2 and 3.3 for pairwise comparison
data).
44
Treatment 1
Visual Surface Area
Treatment 2
Median 1
Visual Surface Area
Median 2
αa
P-value
Dihydrosphingosine
0.658
Control
1.215
0.0084
<0.0001b
Phytosphingosine
1.356
Control
1.215
0.0084
0.9725
Sphingosine
0.684
Control
1.215
0.0084
<0.0001b
Phytosphingosine
1.356
Dihydrosphingosine
0.658
0.0084
<0.0001b
Sphingosine
0.684
Dihydrosphingosine
0.658
0.0084
0.6784
Sphingosine
0.684
Phytosphingosine
1.356
0.0084
<0.0001b
Table 3.2. E. coli pairwise exact Wilcoxon Rank Sum treatment comparisons of the visual surface area.
a
The Bonferroni correction was used to adjust for multiple pairwise comparisons to maintain an overall significance level of 0.05.
b
Significant after adjustment for pairwise multiple comparisons.
45
Treatment 1
Visual Surface Area
Median 1
Treatment 2
Visual Surface Area
Median 2
αa
P-value
Dihydrosphingosine
0.128
Control
0.219
0.0084
0.0001b
Phytosphingosine
0.215
Control
0.219
0.0084
1.0000
Sphingosine
0.135
Control
0.219
0.0084
<0.0001b
Phytosphingosine
0.215
Dihydrosphingosine
0.128
0.0084
<0.0001b
Sphingosine
0.135
Dihydrosphingosine
0.128
0.0084
0.2993
Sphingosine
0.135
Phytosphingosine
0.215
0.0084
<0.0001b
Table 3.3. S. aureus pairwise exact Wilcoxon Rank Sum treatment comparisons of the visual surface area.
a
The Bonferroni correction was used to adjust for multiple pairwise comparisons to maintain an overall significance level of 0.05.
b
Significant after adjustment for pairwise multiple comparisons.
46
47
Figure 3.1. SEM images of E. coli and S. aureus untreated and treated with
sphingoid bases. Sphingoid base treatments were phytosphingosine: C-D;
sphingosine: E-F; dihydrosphingosine: G-H. Sphingoid base-treated E. coli
(left panel) were distorted and their surfaces were concave and rugate, with
many external blebs. Sphingoid base-treated S. aureus (right panel) were also
distorted with some cells appearing concave. Many cells were in various
stages of lysis with compromise of the cell wall and plasma membrane.
Untreated bacteria (A,B) exhibited normal morphology.
48
Figure 3.2. TEM images of E. coli and S. aureus untreated and treated with
sphingoid bases. Sphingoid base treatments were phytosphingosine: C-D;
sphingosine: E-F; dihydrosphingosine: G-H. Size bars are equal to 0.2 µm.
E. coli (left panel) treated with sphingoid bases contained electron dense
intracellular bodies not present in the control bacteria (A) and the remaining
cytoplasm was not uniform, suggesting flocculation of the intracellular
contents. S. aureus (right panel) treated with sphingoid bases also contained
electron dense intracellular inclusion bodies not seen in control samples (B)
with aggregation of the remaining intracellular contents. Cells were also in
various stages of disintegration and lysis with sections of cell wall and cellular
debris visible near damaged cells.
49
Figure 3.3. Association of E. coli and S. aureus lipids with sphingoid bases after
treatment. Densiometry measurements of the carbon present in the
sphingosine standard lanes of TLC chromatograms were used to estimate the
total extracted lipid weight in each of the treated and untreated bacterial
samples as well as lipid-only controls. Percentage of lipid uptake for each
sample was calculated by dividing the total extracted lipid weight by the total
weight of lipid added. Both E. coli and S. aureus took up a large percentage
of the added sphingoid bases relative to controls. Controls included
sphingosine (white bar), phytosphingosine (gray bar), and dihydrosphingosine
(black bar) in media to ensure that they would not sediment without true
bacterial association.
50
CHAPTER 4
ORAL MUCOSAL LIPIDS ARE ANTIBACTERIAL AGAINST
PORPHYROMONAS GINGIVALIS, INDUCE ULTRASTRUCTURAL
DAMAGE, AND ALTER BACTERIAL LIPID AND PROTEIN
COMPOSITIONS
Infection and inflammation in the oral cavity ranges from gingivitis, a mild and
reversible inflammation of the gingiva, to aggressive periodontitis, a chronic
inflammation and associated exaggerated immune response (Berglundh and Donati 2005)
that leads to progressive destruction of the periodontal ligament and alveolar bone.
Dependent upon oral hygiene, socio-economic status, and other environmental, genetic
and metabolic risk factors, periodontitis occurs in just over 47% of the population of the
United States with a prevalence of 8.7, 30.0, and 8.5% for mild, moderate, and severe
periodontitis, respectively (Eke et al. 2012).
P. gingivalis, one of more than 600 bacterial species found in the oral cavity, is
among the most influential of periodontal pathogens; P. gingivalis is more likely to be
found in patients with periodontitis and less likely to be present in healthy individuals.
Furthermore, P. gingivalis shows a strong positive relationship with two parameters
important in the diagnosis of periodontitis: increased sulcular pocket depth and bleeding
upon probing (Hutter et al. 2003, Socransky and Haffajee 1992, Socransky et al. 1998).
This gram-negative, black pigmented, strict anaerobic coccobacillus is recognized as a
late colonizer in the development of oral biofilms (Kolenbrander et al. 2002, Socransky et
al. 1998), where the multitude of virulence factors produced by P. gingivalis contributes
to its pathogenicity (Holt et al. 1999). Additionally, P. gingivalis produces many
proteins, enzymes, and metabolic end products that are important to its survival and
growth within the host because they are active against a broad spectrum of host proteins
and provide mechanisms for evasion of host defenses (Holt et al. 1999).
51
Control of oral bacteria is mediated by a diverse array of specific and non-specific
innate immune factors present in saliva and on mucosal surfaces (Gorr 2009, Gorr 2012).
More than 45 AMPs are grouped into functional families that include cationic peptides,
metal ion chelators, histatins, defensins, bacterial adhesions and agglutinators, and
enzymes directed at the bacterial cell wall. The physiologic concentration of most
salivary AMPs, however, is lower than the effective concentration in vivo (Gorr 2012)
which suggests that there may be additional immune functions within the saliva.
Lipids, although less well known, are also important innate immune molecules
(Bibel et al. 1993, Drake et al. 2008). Saliva contains an array of lipids that include
cholesterol, fatty acids, triglycerides, wax esters, cholesterol esters, and squalene (Brasser
et al. 2011a, Brasser et al. 2011b, Law et al. 1995a, Law et al. 1995b). These lipids
contribute to a variety of cellular and immune-related processes including transport of
fat-soluble antioxidants to and from the mucosal surfaces, the pro- and anti-inflammatory
properties of mucosal surfaces, and the innate antimicrobial activity of mucosal surfaces
(Smith and Thiboutot 2008, Zouboulis 2004, Zouboulis et al. 2008). Sphingoid bases and
short chain fatty acids, of epithelial and sebaceous gland origin, are found within the
saliva, the stratum corneum of the gingiva and hard palate, and the mucosal epithelium.
These sphingoid bases and short chain fatty acids exhibit antimicrobial activity against a
variety of gram-positive and gram-negative bacteria (Bergsson et al. 2001a, Bergsson et
al. 2002, Bibel et al. 1992b, Bibel et al. 1993, Burtenshaw 1942). Recent work suggests
these lipids are also likely involved in innate immune defense against epidermal and
mucosal bacterial infections (Drake et al. 2008, Law et al. 1995b). However, relatively
little is known about the spectrum of lipid activity against oral bacteria or the
mechanisms of action.
In this study, we examine the antimicrobial activity of sphingoid bases:
sphingosine, dihydrosphingosine, and phytosphingosine, and fatty acids: sapienic acid
and lauric acid, commonly found within the oral cavity, against P. gingivalis. We also
52
explore potential mechanisms of action for select lipid-organism combinations and
present their potential as pharmaceuticals to improve therapies for treatment of mucosal
infections and inflammatory disorders.
Materials and Methods
Bacterial species and growth conditions
P. gingivalis strain 381 ATCC BAA 1703 was cultured in TSB (Difco
Laboratories, Detroit, MI) supplemented with vitamin K1 and hemin (Sigma Chemical
Co., St. Louis, MO) and incubated at 37ºC in an anaerobic chamber (Coy Laboratory
Products Inc., Grass Lake, MI) containing an atmosphere of 85% N2, 10% H2, and 5%
CO2. Unless otherwise noted, we transferred cells to fresh medium and grew them
overnight before adjusting to contain 1 × 108 CFU/ml (0.108 O.D., 600 nm, Spectronic
20D+, Thermo Fisher Scientific, Inc., Waltham, MA) and then diluting to a concentration
of 1 × 107 CFU/ml. Unless otherwise noted, controls for all assays included medium
only (sterility control), P. gingivalis treated with 0.14 M NaCl (negative treatment control
and positive growth control), and SMAP28 (positive control).
Preparation of lipids
Phytosphingosine, sphingosine, dihydrosphingosine, and lauric acid were
obtained from Sigma Chemical Company (St. Louis, MO). Sapienic acid was obtained
from Matreya Inc. (Pleasant Gap, PA). Stock solutions were prepared as described in
Chapter 2 and lipids were diluted to the desired concentration using 0.14 M NaCl.
Antimicrobial assay
Using broth microdilution assays, we determined the MIC for each bacteria-lipid
combination (Brogden et al. 2001). We serially diluted lipids in 0.14 M NaCl (500 to 1
g/ml) in microtiter plates (Immunolon 1 microtiter plates, Thomas Scientific,
Swedesboro, NJ) and added P. gingivalis at a concentration of 1 × 107 CFU/ml. After
53
incubation for five days as described above, we read the OD (λ = 600 nm) of bacterial
growth in a spectrophotometer (Spectromax Microplate Reader, Molecular Devices
Corp., Sunnyvale, CA) and determined the MIC. MBCs were determined by plating
bacteria from the completed broth microdilution assays onto CDC formulation anaerobic
5% sheep blood agar plates (Remel, Lenexa, KS). We incubated plates for seven days as
described above before examination of the plates for the presence of CFU.
SMAP28 was included in this study as a positive control to show that the
microdilution assays were set up properly and MICs were accurate and within previously
reported ranges (Bratt et al. 2010b, Weistroffer et al. 2008). SMAP28 was prepared as
described in Chapter 2 and suspended in 0.14 M NaCl for all assays.
Kill kinetics
Using the spiral plating method (Drake et al. 1994), we assessed kill kinetics for
each lipid against P. gingivalis. For this, we prepared a 1 × 107 CFU/ml suspension of P.
gingivalis, divided this suspension into tubes for each treatment, and added either 0.14 M
NaCl or each of the lipids at a concentration equivalent to 10X the MIC determined in the
broth microdilution assays. At time intervals of 0, 0.5, 1, 2, 3, 4, 6, 8, and 24 hours, we
serially diluted one-ml samples from each treatment into 0.14 M NaCl and plated the
diluted samples onto CDC formulation anaerobic 5% sheep blood agar plates (Remel,
Lenexa, KS) using an Autoplate 4000 Automated Spiral Plater (Advanced Instruments,
Inc. Norwood, MA). After incubating for seven days we counted the CFU and calculated
concentrations.
Ultrastructural analyses of lipid-exposed bacterial cells
Broth cultures of P. gingivalis were adjusted to 1 × 107 CFU/ml in growth media
as described above, and treated with 80 µg/ml phytosphingosine, 586 µg/ml sapienic
acid, 50 µg/ml SMAP28, or 0.14 M NaCl for one hour. To visualize cells in various
54
stages of death, we based incubation times on kill kinetics so that each suspension
contained both viable (<50%) and non-viable (≥50%) cells.
For examination by TEM, treated P. gingivalis were fixed using 2.5%
glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for one hour in an ice bath,
and washed twice in 0.1 M sodium cacodylate buffer (pH 7.4) for 20 minutes. We then
pelleted the bacteria by centrifugation, suspended the cells in warm 0.9% agarose in 0.1
M sodium cacodylate buffer, pH 7.4, and allowed the agarose to congeal before dicing it
into one-mm cubes. After two washes in 0.1 M sodium cacodylate buffer, pH 7.4, for 20
minutes, we treated the cubes with 1% osmium tetroxide for one hour, washed them
again in 0.1 M sodium cacodylate buffer, and then dehydrated the cubes in a series of
30%, 50%, 70%, 95%, and absolute ethanol solutions. After clearing in propylene oxide,
we infiltrated the cubes with a propylene oxide-Epon mixture (1:1), embedded them in
Epon, and polymerized at 60oC for 48 hours. Finally, we cut ultrathin sections from each
cube, placed sections on formvar-coated nickel grids, and stained with 5% uranyl acetate
and Reynold’s lead citrate. We examined samples for intracellular damage using a JEOL
JEM-1230 TEM (JEOL USA, Inc., Peabody, MA USA).
For examination by SEM, treated or untreated P. gingivalis were layered on a
nucleopore membrane (SPI Supplies, West Chester, PA), fixed with 2.5% glutaraldehyde
in 0.1 M sodium cacodylate buffer (pH 7.4) for one hour in an ice bath, and washed twice
in 0.1 M sodium cacodylate buffer (pH 7.4) for four minutes. We then further fixed
samples with 1% osmium tetroxide for 30 minutes, washed them twice in double distilled
water, and then dehydrated them in a series of 25%, 50%, 75%, 95%, and absolute
ethanol solutions followed by hexamethyldisilizane. After mounting the membranes
containing bacteria onto stubs, we sputter coated them with gold and palladium, and
examined each sample for surface damage using a Hitachi S-4800 field emission SEM
(Hitachi High-Technologies Canada, Inc., Toronto, Ontario Canada).
55
Lipid analysis
Broth cultures of P. gingivalis were incubated with each of sphingosine,
dihydrosphingosine, phytosphingosine, sapienic acid, lauric acid, and 0.14 M NaCl at 500
µg/ml (total volume of each treatment was 5 ml) for 1.5 hours at 37°C. After treatment
with lipids, we divided each sample and processed half for lipid analysis and half for
protein analysis (next section). Before pelleting by centrifugation, bacteria were killed by
adding 0.05% sodium azide. After freezing these whole cell pellets at -80°C, we
lyophilized the bacteria and extracted the lipids using a previously described method
(Wertz et al. 1987) consisting of successive extractions of chloroform:methanol mixtures
(2:1, 1:1, and 1:2) at room temperature. Extracted lipids were recovered by evaporation
of the solvent under a stream of nitrogen. To purify the samples, we redissolved each
sample in five ml chloroform:methanol (2:1) and washed the solution with one ml 2 M
potassium chloride (20% by volume) to remove salts and other water soluble materials
(Folch et al. 1957). The resulting upper phase was discarded and the lower phase,
containing purified lipids, was again dried under nitrogen. The dried lipids were
reconstituted in chloroform:methanol, 2:1 at a concentration of 10 mg/ml. Additional
controls included suspensions of each treatment lipid in bacterial sterile growth medium
followed by centrifugation and resuspension in chloroform:methanol (2:1) to test the
ability of each lipid to sediment or adhere to the tube, which would cause false positive
results.
The lipids from each treatment and control were separated by QTLC as previously
described (Wertz and Downing 1989). We obtained glass-backed plates coated with a
500 µm thickness of silica G gel (Alltech Associates, Deerfield, IL) and prepared the
plates by washing with chloroform:methanol (2:1) to remove organic contaminants.
Plates were then air-dried and activated in a 110°C oven. After dividing the silica gel G
plates into six-mm wide lanes, we spotted total extracted lipids from each sample onto
the lanes and developed these chromatograms differentially for each lipid class.
56
Chromatograms for separation of sphingoid bases were developed in
chloroform:methanol:water (40:10:1). Sphingosine served as a standard for
quantification (Weerheim and Ponec 2001). For separation of fatty acids, chromatograms
were developed in three sequential solvent mixtures: 1) n-hexane; 2) toluene; and 3)
hexane:ethyl-ether:acetic acid (70:30:1). A standard containing squalene, cholesterol
esters, wax esters, triglycerides, fatty acids, and cholesterol was used to identify
migration of the fatty acids. For development of chromatograms, we sprayed each plate
with 50% sulfuric acid and charred the lipid bands by heating slowly to 220°C on a
hotplate. Digital images were obtained using a Hewlett-Packard Scanjet 3500c and
analyzed using TNIMAGE (Thomas Nelson, Bethesda, MD) in strip densiometry mode
to estimate the total extracted lipid weight in each of the treated and untreated bacterial
samples as well as controls. To calculate the percentage of lipid uptake for each sample
we divided the total extracted lipid weight by the total weight of lipid added to each
sample. Because P. gingivalis plasma membrane naturally contains dihydrosphingosine
(Mun et al. 2007, Nichols et al. 2004, Nichols 1998, Nichols et al. 2006), total sphingoid
base lipids were normalized by subtracting the total sphingoid base weight present in the
untreated P. gingivalis controls.
Protein analyses
For analysis by reducing-SDS-PAGE (sodium dodecyl sulfate polyacrylamide gel
electrophoresis), lipid-treated and untreated P. gingivalis samples (using the remaining
half of samples processed for lipid analysis), suspended in SDS (sodium dodecyl sulfate)
reducing sample buffer, were sonicated in a bath sonicator five times at three minutes
each time, cooling on ice between each sonication event, before boiling for eight minutes.
After denaturing the proteins, we loaded the samples onto a NuPage 4-12% BisTris 1.5
mm gel (Life Technologies, Grand Island, NY) and separated the protein fractions using
the XCell SureLockTM Mini-Cell Electrophoresis System (Invitrogen, Carlsbad, CA) in a
57
buffer system of NuPage 1X MOPS SDS running buffer and 0.1% NuPage antioxidant
(Life Technologies, Grand Island, NY). We used Novex Sharp Protein Standards as size
markers (Life Technologies, Grand Island, NY).
Protein bands were then either visualized using the GenScript eStain Protein
Staining System (Piscataway, NJ), or transferred onto polyvinylidene difluoride (PVDF)
membranes (Life Technologies, Grand Island, NY) for Western blot analysis using the
XCell II Blot Module Western blot system (Invitrogen, Carlsbad, CA) in a buffer of
NuPage 1X transfer buffer with 0.1% antioxidant (Life Technologies, Grand Island, NY).
After transfer of proteins to the PVDF membrane, we visualized proteins using a 0.1%
Coomassie Brilliant Blue R-250 stain (Sigma Chemical) in 40% methanol and 10% acetic
acid, followed by destaining in methanol:acetic acid:water solutions (40:10:50 followed
by 90:5:5). Bands of interest were excised and sequenced (Protein Facility, Iowa State
University, Ames, IA) by the Edman N-terminus degradation process and BLAST
searches of the National Center for Biotechnology Information (NCBI) P. gingivalis
protein database identified upregulated protein bands of interest.
In addition, we used two-dimensional difference in-gel electrophoresis (2DDIGE) (Applied Biomics, Hayward, CA) to compare proteins present in sapienic acidtreated and untreated P. gingivalis. For 2D-DIGE, treated and untreated samples were
labeled with different fluorescent dyes, mixed, and then separated by isoelectric point
followed by molecular weight separation. From the resulting gels, we chose 16 spots
indicative of upregulated proteins in the sapienic acid-treated sample for sequencing by
mass spectroscopy. Sequences were identified by an NCBInr BLAST search of the P.
gingivalis protein database.
Statistical analyses
Preliminary evaluation of MIC, MBC, and kill kinetics data using the ShapiroWilk procedure provided strong evidence of departure from normality; consequently
58
nonparametric procedures were used throughout. The Kruskal-Wallis test was employed
to detect treatment differences in MIC and MBC distribution; the adaptation of the Tukey
method due to Conover (Conover 1999) was used to adjust for multiple pairwise
comparisons of lipid treatment groups in conjunction with an overall 5% level of
significance.
Two summary measures of kill kinetics were computed for comparison of
longitudinal data between treatment groups. Trapezoidal AUC was used as a summary
measure of bacterial viability over the treatment time course (Dawson and Siegler 1996,
Ghosh et al. 1973) where larger AUC values correspond to greater viability.
Comparisons were made with and without the inclusion of AUCs from the control
sample. A second summary measure of kill kinetics over time considered was time to
zero, defined as the first time point at which total bacterial counts reached zero (complete
killing). Because samples sizes were modest, the overall test for treatment differences for
these two outcomes was conducted using exact Kruskal-Wallis tests. Pair-wise
comparisons were made using exact Wilcoxon Rank Sum tests with Bonferroni
correction for multiple comparisons, again in conjunction with an experiment-wise Type
I error level of 5%. Note that, for certain of these longitudinal assays (i.e. from a given
vial), none of the bacterial counts in the series reached zero. In such instances, the value
of the corresponding time to zero was assigned the highest rank for purposes of analysis.
If several such instances occurred in a given analysis, ties corresponding to the highest
rank were assigned.
Results
All lipids exhibited antimicrobial activity against P. gingivalis with variability in
activity across lipids, ranging in MIC from 0.2 – 125.0 µg/ml and in MBCs from 0.3 –
218.8 µg/ml (Table 4.1). Distribution of both MIC and MBC values differed among the
59
treatment groups (p < 0.0001) and all ten pairwise comparisons were significantly
different for MICs (p values <0.0001-0.0014) and MBCs (p values <0.0001 – 0.0002).
Sapienic acid rapidly killed P. gingivalis, with complete death occurring before
the first sampling time of six minutes (Figure 4.1). The remaining lipid treatments
greatly reduced the bacterial count within six minutes with complete killing occurring in
all most instances within 30 minutes. Phytosphingosine had the longest time to zero at
one hour. Following adjustment for multiple comparisons, significant differences in time
to zero were identified between sapienic acid and phytosphinogosine, as well as between
these two lipids and each of the other three treatments (Table 4.2). Using these analyses,
no significant difference was found between dihydrosphingosine, sphingosine, and lauric
acid, as all three treatments had a median time to zero of 30 minutes. However,
trapezoidal AUC, calculated for each lipid treatment over the time interval 0.1 hours to
24 hours (Table 4.3) highlights the differences across treatments. After Bonferonni
adjustment (adjusted α = 0.033) for fifteen comparisons, the outcome for each treatment
was found to significantly differ from that of each of the others (p < 0.0022) over this
time period.
SEM micrographs demonstrated that P. gingivalis cells treated with
phytosphingosine (Figure 4.2, B1-2) or sapienic acid (Figure 4.2, C1-2) showed various
stages of lysis. Cellular debris and detached pieces of membrane lay adjacent to the cells.
Many cells were distorted with a concave and rugate appearance and loss of cellular
content. In addition, the cells were more closely aggregated and increased numbers of
external blebs (relative to controls) were present on and around the bacteria. Similar to
lipid-treated bacteria, SMAP28-treated P. gingivalis cells (Figure 4.2, D1-2) were also
distorted with concave and rugate morphology and were in various stages of lysis with
loss of intracellular content. Untreated P. gingivalis (Figure 4.2, A1-2) cells exhibited an
external structure typical of healthy gram-negative coccobacilli (Holt et al. 1999) with
multiple blebs present on the cell surface (Figure 4.2, A2).
60
Examination of untreated P. gingivalis thin sections by TEM (Figure 4.3, A1-A3)
revealed typical gram-negative morphology (Mansheim and Coleman 1980, Mayrand and
Holt 1988, Parent et al. 1986) and internal structures were visible. All lipid-treated and
SMAP28-treated cells, however, exhibited intracellular damage. Detached membrane
was lying adjacent to damaged cells and increased numbers of blebs (relative to controls)
were present on and around the cells. Phytosphingosine (Figure 4.3, B1-4) and SMAP28
(Figure 4.3, D1-4) treatment induced separation of the outer membrane from the
cytoplasmic membrane. Plasma membranes were compromised, with leakage of cellular
contents. Both treatments also caused a loss of distinct nucleoid and ribosomal regions in
many cells and a decrease in the electron density of the cytoplasmic contents. Treatment
with sapienic acid (Figure 4.3, C1-4) induced a different type of membrane disruption.
Many sapienic acid-treated cells exhibited a bunching or “scrubbing” of the outer
membrane. Pieces of the cell wall/membrane complex were missing in many cells and
loose membrane pieces were lying adjacent to damaged cells, resulting in leakage of
cellular contents.
Chromatographic separation of total lipid extracts from fatty acid or sphingoid
base-treated P. gingivalis confirmed the presence of considerable amounts of treatment
lipid in every sample relative to untreated P. gingivalis controls (Figure 4.4). P.
gingivalis retained 30-55% of the treatment lipids added to each sample, indicating
association of both fatty acids and sphingoid bases with P. gingivalis lipids. Uptake of
treatment lipids varied across treatments with fatty acids showing more association with
bacterial lipids than sphingoid bases.
P. gingivalis protein expression also changed with lipid treatment. Protein
analysis by SDS-PAGE revealed differential banding patterns between untreated and
lipid-treated P. gingivalis samples. The most striking differences were seen with sapienic
acid treatment (Figure 4.5). Further analysis of sapienic acid-treated P. gingivalis
through Western blot and 2D-DIGE demonstrated the differential expression of many
61
proteins relative to an untreated sample. Upon sequencing of 16 upregulated protein
spots from the 2D-DIGE gel (Figure 4.6), and seven bands from the Western blot, we
found proteins involved in biosynthesis of bacterial lipids, metabolism and energy
production, metabolism in diverse environments, amino acid biosynthesis, acquisition of
peptides, degradation of polypeptides, cell adhesion, and virulence (Table 4.4).
Discussion
In this study we report for the first time, to our knowledge, that lipids endogenous
to saliva and oral mucosa are antimicrobial for P. gingivalis and induce novel
ultrastructural damage. Our results are in agreement with growing evidence that fatty
acids and sphingoid bases differentially kill bacteria in a dose-dependent manner and
induce cellular damage. For example, E. coli and S. aureus treated with sphingosine,
phytosphingosine, or dihydrosphingosine exhibit extensive and differential intracellular
and extracellular damage (Fischer et al. 2013). Bibel and colleagues (Bibel et al. 1993)
also showed that sphinganine (e.g. dihydrosphingosine) treatment of S. aureus results in
ultrastructural damage similar to antibiotic treatment, including lesions of the cell wall,
membrane evaginations, and leakage. In addition, treatment of Helicobacter pylori with
oleic or linoleic acid exhibits altered morphology with disruption of cellular membranes
and cell lysis (Khulusi et al. 1995).
Our work indicates that there may be different mechanisms involved for the
activity of different lipids. Antimicrobial activity, the percentage of lipid retained by P.
gingivalis, and ultrastructural damage are all dependent upon the specific lipid treatment.
These data, combined with our observation that fatty acids and sphingoid bases exhibit
differential activity across bacterial species (Fischer et al. 2012), lead us to believe that
the antimicrobial activity of fatty acids and sphingoid bases is a specific interaction that
depends upon characteristics of both the bacterium and a particular lipid. We propose
that mechanisms for the antimicrobial activity of fatty acids and sphingoid bases against
62
bacteria fit within four broad pathways: 1) membrane disruption by detergent activity; 2)
incorporation of lipids into the bacterial plasma membrane; 3) transport of lipids across
the bacterial membrane into the cytosol; and 4) specific interactions between lipids and
protein components of the bacterial membrane. Potential end results of fatty acid
treatment have been reviewed (Desbois and Smith 2010) and include creation of pores in
the bacterial cell, alteration of the cellular membrane, lysis of the cell, and disruption of
various cellular processes either by interference of spatial arrangement or by direct
binding to proteins.
The main site of lipid activity against P. gingivalis is likely the bacterial plasma
membrane, possibly by incorporation of lipids into the membrane. Our results show that
both fatty acids and sphingoid bases are retained by P. gingivalis after treatment. In
addition, destruction of the membrane is evident in TEM micrographs. This is similar to
activity seen in other organisms following fatty acid and sphingoid base treatment. S.
aureus treated with capric acid exhibits damage to the membrane but not the cell wall
(Bergsson et al. 2001a). Furthermore, L-forms of S. aureus (lacking cell walls) are
relatively resistant to the lethal effects of dihydrosphingosine, suggesting that the plasma
membrane is necessary for activity (Bibel et al. 1993). H. pylori treated by two fatty
acids, linoleic acid and oleic acid, also exhibits membrane destruction and both fatty
acids incorporate into the plasma membrane, altering the phospholipid composition of H.
pylori (Khulusi et al. 1995).
Activity of fatty acids and sphingoid bases are also likely dependent upon the
specific phospholipid composition of the bacterial plasma membrane. In this study we
show that sphingoid bases are more active against P. gingivalis than a variety of other
gram-positive and gram-negative bacteria previously examined (Fischer et al. 2012). P.
gingivalis contains several classes of novel phospholipids and branched lipids (Nichols et
al. 2004, Nichols 1998, Nichols et al. 2006) including phosphorylated dihydroceramides
(a source of dihydrosphingosine). Because the P. gingivalis bacterial membrane contains
63
sphingolipids, sphingoid bases may be more likely to either incorporate into the bacterial
membrane or pass through the membrane. It is also possible that P. gingivalis may
attempt to either utilize sphingoid bases for building its unique phospholipids or as an
energy source. In our 2D-DIGE analysis of sapienic acid-treated P. gingivalis, we found
upregulation of two key regulators of lipid metabolism, involved in catalyzing the
condensation reaction of fatty acid biosynthesis: 3-oxoacyl-synthase-2 and 2-oxoacylsynthase-3. Increasing production of fatty acids could serve several purposes: 1)
increasing phospholipid production to repair damaged bacterial membranes; 2) utilization
of introduced fatty acids or sphingoid bases for phospholipid production (which may or
may not be harmful); 3) competition with harmful sphingoid bases that could insert into
the plasma membrane.
Activity at the bacterial membrane may also depend upon the structure and shape
of the treatment lipids. Several lipid characteristics important for activity include:
hydrophobicity, number, placement, and orientation of double bonds (Kabara et al.
1972a, Saito et al. 1984), and in fatty acids, the length of the carbon chain (Kabara et al.
1972a, Willett and Morse 1966, Zheng et al. 2005) and the –OH group (Zheng et al.
2005). Studies indicate that fatty acids with cis-double bonds are more active than fatty
acids with trans-double bonds (Galbraith et al. 1971, Kabara et al. 1972a). A cis-bonded
lipid would likely cause a fluidizing effect upon insertion into a bacterial plasma
membrane.
Finally, we show that sapienic acid induces upregulation of a unique set of
proteins that may provide clues to specific mechanisms of action. In our Western blot
and 2D-DIGE analysis of sapienic acid-treated P. gingivalis, we found upregulated
proteins important in various cellular processes including glycolysis, amino acid
metabolic processes, microbial metabolism in diverse environments, acquisition and
degradation of polypeptides, adhesion, and other virulence factors. P. gingivalis exhibits
several unique stress responses, dependent upon the type of stressor. Heat stress (Amano
64
et al. 1994, Bonass et al. 2000, Lopatin et al. 1999, Lu and McBride 1994, Murakami et
al. 2004, Percival et al. 1999, Shelburne et al. 2005, Vayssier et al. 1994), O2 oxidative
stress (Meuric et al. 2008, Shelburne et al. 2005, Vayssier et al. 1994), H2O2 oxidative
stress (Meuric et al. 2008, Shelburne et al. 2005, Vanterpool et al. 2010), pH stress (Lu
and McBride 1994, Vayssier et al. 1994), heme limitation (Dashper et al. 2009), ethanol
stress (Lu and McBride 1994), and response to contact with epithelial cells (Hosogi and
Duncan 2005) all induce extensive and unique responses in P. gingivalis with very little
overlap. These well-documented stress responses have very little in common with the
response induced by sapienic acid treatment (Appendix A).
All these data combined suggest that with sapienic acid, there may be a quick
two-step process leading to antimicrobial activity that is dependent upon time and
sapienic acid concentration. As P. gingivalis cells are exposed to sapienic acid they
begin taking up large amounts of the lipid, become stressed and quickly mount a response
by adjusting protein activity, as evidenced by the differential protein profiles and the
upregulation of several components important for microbial metabolism in diverse
environments. It is possible, however, that as a critical point (time and/or lipid
concentration) is reached, rescue attempts fail and these cells succumb to lysis. Further
analysis of the metabolic consequences of sapienic acid treatment on P. gingivalis will be
necessary to confirm this and will possibly be the subject of future studies.
The “self-disinfecting” properties of the skin have been recognized since 1942
when Burtenshaw described skin lipids that were active against a number of bacteria
(Burtenshaw 1942). Recent studies indicate that fatty acids and sphingoid bases function
as innate immune molecules on the skin (Brogden et al. 2011, Drake et al. 2008), oral
mucosa (Bratt et al. 2010a), and in other body fluids such as breast milk (Field 2005,
Hosea Blewett et al. 2008) and sebum (Wille and Kydonieus 2003). In addition, lipid
deficiencies or imbalances in lipid ratios are associated with several diseases. For
example, both deficient hexadecanoic acid production (Takigawa et al. 2005) and
65
decreased levels of sphingosine (Arikawa et al. 2002) are associated with atopic
dermatitis and subsequent increase in S. aureus skin colonization within otherwise
healthy individuals. In addition, cystic fibrosis is linked with deficient fatty acid
production (Freedman et al. 2004, Strandvik et al. 2001). In another study, failure to
clear skin infections of S. aureus or Streptococcus pyogenes within innate
immunodeficient mice was linked to mutation of an enzyme necessary for palmitic and
oleic acid production (Georgel et al. 2005). Based on this information, it is possible that
imbalances in lipid ratios or defective production of certain lipids could be responsible
for other skin and oral diseases. It becomes reasonable then to speculate that topical
application of endogenous lipid formulations could potentially supplement the natural
immune function of lipids on skin and other mucosal surfaces (Thormar and Hilmarsson
2007).
With the increasing resistance of bacteria to many available antibiotic treatments
(Thormar and Hilmarsson 2007) it becomes more important to look for alternative
treatments. Undecylenic acid has been used in over-the-counter antifungal preparations
for several years (Anonymous 2002, Shapiro and Rothman 1983). Hydrogels containing
lipid suspensions are also appearing in literature as topical treatments for a variety of
viruses and bacteria (Neyts et al. 2000, Thormar et al. 1999) and have been used in mice
with no apparent irritation or toxic side effects (Neyts et al. 2000). Clinical use of
endogenous lipids would have several advantages over other antibiotic treatments.
Drake, et al. (Drake et al. 2008) point out that because they are normal occupants of the
skin [and oral mucosa] lipids are likely to be less irritating. In addition, because of their
evolution with the potential pathogens of skin and oral mucosa it is more unlikely that
these pathogens will readily develop resistance to treatment (Drake et al. 2008).
Additionally, fatty acids and sphingoid bases used in our studies were active within
normal physiologic ranges (4.0 - 13.2 µg/ml for total fatty acids and 0.5 – 5.0 µg/ml for
66
free sphingoid bases) (Brasser et al. 2011a, Brasser et al. 2011b) and would therefore be
effective in tolerable concentrations.
Crucial to the development of formulations that would stimulate the natural innate
function is a better understanding of the spectrum of fatty acid and sphingoid base
activities and mechanisms of action. Knowledge of mechanisms behind the antimicrobial
activity of antibacterial lipids is sparse and these data contribute to the available
information.
67
MIC Meana
(Median)
MBC Meana
(Median)
Sphingosine
0.2 ± 0.8
(0.2)
0.3 ± 0.0
(0.3)
Phytosphingosine
0.8 ± 0.3
(0.8)
1.0 ± 0.0
(1.0)
Dihydrosphingosine
0.4 ± 0.2
(0.4)
0.5 ± 0.2
(0.6)
Sapienic acid
58.6 ± 11.0
(58.6)
62.5 ± 0.0
(62.5)
Lauric acid
125.0 ± 0.0
(125.0)
218.8 ± 57.9
(250.0)
5.0 ± 0.0
(5.0)
20.0 ± 0.0
(20.0)
Treatment
SMAP-28
Table 4.1. Minimum lipid concentrations required to inhibit or kill P. gingivalis.
a
Mean ± standard deviation (median); n = 8 per treatment.
Note: Overall comparison of treatments by Kruskal-Wallis showed significant
distribution differences across treatment groups for both MICs and MBCs (p < 0.0001
for each). Pairwise comparisons of all MICs and MBCs (µg/ml ± SD) for all lipid
treatments against P. gingivalis showed significantly different outcomes for all
treatments at a 5% level of statistical significance. SMAP28 was used as a positive
control to show that the microdilution assays were set up properly and MICs/MBCs
were accurate and within previously reported ranges. SMAP28 results were not
included in the statistical analyses.
68
Treatment 1
Median
(hours)
Treatment 2
Median
(hours)
P-values
Dihydrosphingosine
Dihydrosphingosine
Dihydrosphingosine
Dihydrosphingosine
0.5
0.5
0.5
0.5
Lauric Acid
Phytosphingosine
Sapienic Acid
Sphingosine
0.5
1.0
0.1
0.5
1.0000
0.0022a
0.0022a
1.0000
Lauric Acid
0.5
Phytosphingosine
1.0
0.0022a
Lauric Acid
Lauric Acid
Phytosphingosine
0.5
0.5
1.0
Sapienic Acid
Sphingosine
Sapienic Acid
0.1
0.5
0.1
0.0022a
1.0000
0.0022a
Phytosphingosine
Sapienic Acid
1.0
0.1
Sphingosine
Sphingosine
0.5
0.5
0.0022a
0.0022a
Table 4.2. Pairwise comparisons of the time required to kill P. gingivalis by each of
the lipid treatments.
a
Indicates significance after adjustment for multiple comparisons.
Note: Pairwise comparisons were Bonferonni adjusted for ten comparisons (α = 0.005) at
a 5% significance level.
69
Treatment
Mean
SD
Median
IQR
Min
Max
Control
Dihydrosphingosine
Lauric Acid
Phytosphingosine
574.05
3.22
1.73
5.34
1.19
0.03
0.03
0.30
573.97
3.23
1.73
5.25
1.68
0.03
0.04
0.51
572.65
3.16
1.70
5.06
575.72
3.25
1.76
5.76
Sapienic Acid
0.00
0.00
0.00
0.00
0.00
0.00
Sphingosine
1.28
0.09
1.27
0.05
1.19
1.45
Table 4.3. AUC analysis of kill kinetics.
Note: Trapezoidal area under the kill kinetics curve was calculated as a summary
measure of P. gingivalis viability over the treatment time course of 0.1 – 24 hours.
For each treatment n = 6 replicates. An exact Kruskal-Wallis test followed by
pairwise comparisons showed that after Bonferroni adjustment (α = 0.0033) the
trapezoidal area distribution is significantly different for all treatments (p = 0.0022 for
all comparisons).
Protein (Identification source)
Gene
Accession#
Sequence length (aa)
MW (Daltons)
Function
Biological Process
3-oxoacyl-[acyl-carrier-protein] synthase
2 (2D-DIGE)
fabF
gi|34541387
418
44491.4
Transferase
Fatty acid biosynthesis; fatty acid elongation
3-oxoacyl-[acyl-carrier-protein] synthase
3 (KASIII) (2D-DIGE)
fabH
FABH_PORGI
335
37174.4
Transferase
Fatty acid biosynthesis, elongation
NAD-dependent glutamate
dehydrogenase (GDH) (2D-DIGE & WB)
gdh
gi|334146994; AAA50985
437
48218.8
Oxidoreductase
Cellular amino acid metabolic processes (R, P, A, D, and
E); nitrogen metabolism, virulence (cytotoxic byproducts); glutamate energy metabolism; degradation of
amino acids (energy source)
Glyceraldehyde 3-phosphate
dehydrogenase, type I (2D-DIGE)
gapA
gi|34541701
336
35992.4
Oxidoreductase/NAD binding
Microbial metabolism in diverse environments;
glycolysis/gluconeogenesis; biosynthesis of secondary
metabolites
Table 4.4. Identification of P. gingivalis upregulated proteins upon treatment with sapienic acid.
Note: Identification was completed by 2D-DIGE separation followed by sequencing by mass spectroscopy or by Western blot (WB)
followed by sequencing via n-terminus degradation.
70
Protein (Identification source)
Gene
Accession#
Sequence length (aa)
MW (Daltons)
Function
Biological Process
Phosphoserine aminotransferase (2DDIGE)
serC
gi|334147974
360
40090.6
Aminotransferase
Microbial metabolism in diverse environments; methane
metabolism; amino acid metabolism (G, S, and T);
amino acid biosynthesis (S)
Arginine-specific cysteine proteinase
(RGP-1; RgpA; Gingipain A) (2D-DIGE)
rgpA; prtT
P28784
991
108713.3
Virulence
Acquisition of peptides
Metabolism; protein processing
Adhesion
Arginine-specific cysteine proteinase
(RGP-2; RgpB; Gingipain B) (2D-DIGE
& WB)
rgpB
gi|1814394
736
80952.1
Virulence
Acquisition of peptides
Metabolism; protein processing
Adhesion
Pg-II fimbriae (2D-DIGE)
fimA
gi|22255316
370
39307.8
Virulence
Adhesion
Lysine-specific cysteine protease (Kgp;
Lys-gingipain;) (2D-DIGE & WB)
kgp
Q51817.1
1732
40135.6
Degradation of polypeptides
Table 4.4. Continued
71
Protein (Identification source)
Gene
Accession#
Sequence length (aa)
MW (Daltons)
Function
Biological Process
Hemagglutinin-like protein (2D-DIGE)
gi|34540264
348
39313.4
Adhesion
Kgp/hemagglutinin (WB)
kgp
AAB49691; AAS68176
1358
Degradation of polypeptides
Adhesion
Glycerate dehydrogenase (WB)
hprA
YP-004509887; GI:333804114
317
Microbial metabolism in diverse environments
Biosynthesis of secondary metabolites
Amino acid metabolism (G, S, T)
Table 4.4. Continued
72
73
Figure 4.1. Kill kinetics for all lipid treatments against P. gingivalis. Geometric mean
of n = 6 is shown for each data point. Error bars represent the SEM; where
error bars are not evident, the SEM was zero. All treatments were started at a
CFU equal to the control; therefore, time zero is equal to that of the control
before the addition of treatment. Where no bacteria were recovered, +1 was
added to zero values before log transformation of the data.
74
Figure 4.2. SEM micrographs showing the effects of sphingoid base and fatty acid
treatment on P. gingivalis. Untreated cells (A1-2) exhibit morphology
typical of P. gingivalis gram-negative coccobacilli. One-hour treatments of P.
gingivalis with phytosphingosine (B1-2), sapienic acid (C1-2), or SMAP28
(D1-2) resulted in evidence of cellular distortion relative to the untreated
bacterium including concave and rugate cells, closer aggregation of cells,
and/or lysis, with detached pieces of membrane lying adjacent to the cells.
75
-
Figure 4.3. TEM micrographs showing the effects of sphingoid base and fatty acid
treatment on P. gingivalis. Untreated cells (A1-3) exhibited typical gramnegative coccobacillus morphology with outer membrane (OM), capsule (C),
periplasmic space (PS), peptidoglycan (PG), cellular membrane (CM), a
distinct nucleoid (N) and ribosomal (R) regions, and outer membrane vesicles
(V) . One-hour treatments of P. gingivalis with phytosphingosine (B1-4),
sapienic acid (C1-4), or SMAP28 (D1-4) resulted in cellular distortion relative
to untreated bacteria. Evidence of ultrastructural damage is indicated by the
colored arrows: separation of the outer membrane from the cytoplasmic
membrane (red); missing pieces of membrane (black); leakage of cytoplasmic
contents (white); and detached membrane lying adjacent to cells (yellow).
76
Figure 4.4. Association of antimicrobial lipids with P. gingivalis lipids after
treatment. Densiometry measurements of the carbon present in the
sphingosine and fatty acid standard lanes were used to estimate the total
extracted lipid weight in each of the treated and untreated bacterial samples as
well as controls. Percentage of lipid uptake by P. gingivalis was calculated by
dividing the total extracted lipid weight by the total weight of lipid added to
each sample. Because P. gingivalis membranes naturally contain
dihydrosphingosine, sphingoid base calculations were normalized (indicated
by an asterisk) by subtracting the total sphingoid base present in untreated
samples. Controls included the same concentration of lipids in media,
processed along with samples to test the abilitity of the lipids to adhere to the
sides of the tube or pellet down with the bacteria.
77
Figure 4.5. SDS-PAGE separation of proteins in untreated and sapienic acid-treated
P. gingivalis. Untreated (Pg381) and sapienic acid-treated P. gingivalis (SA)
proteins were separated by SDS-PAGE and visualized using Coomassie Blue
stain. Molecular weight markers (MWM) are Novex Sharp Protein Standards.
78
Figure 4.6. 2D-DIGE gel showing P. gingivalis protein differences in untreated and
sapienic acid-treated samples. Red spots indicate upregulation of proteins in
treated samples and green spots indicate downregulation of proteins, relative
to the control sample. Yellow spots indicate colocalization, where the same
proteins were present in both samples. Sixteen spots (red arrows) were
chosen for sequencing by mass spectroscopy.
79
CHAPTER 5
ORAL MUCOSAL LIPID CYTOTOXICITY: A STUDY USING
DENDRITIC CELLS
The oral cavity contains an array of lipids that includes cholesterol, fatty acids,
triglycerides, wax esters, cholesterol esters, and squalene (Brasser et al. 2011a, Brasser et
al. 2011b, Law et al. 1995a, Law et al. 1995b). These lipids contribute to a variety of
cellular and immune-related processes including transport of fat-soluble antioxidants to
and from the mucosal surfaces, the pro- and anti-inflammatory properties of mucosal
surfaces, and the innate antimicrobial activity of mucosal surfaces (Smith and Thiboutot
2008, Zouboulis 2004, Zouboulis et al. 2008). Sphingoid bases and short chain fatty
acids of epidermal and sebaceous gland origin are found within the saliva, the stratum
corneum of the gingiva and hard palate, and the mucosal epithelium. These sphingoid
bases and short chain fatty acids exhibit antimicrobial activity against a variety of grampositive and gram-negative bacteria (Bergsson et al. 2001a, Bergsson et al. 2002, Bibel et
al. 1992b, Bibel et al. 1993, Burtenshaw 1942). Recent work suggests these lipids are
also involved in innate immune defense against epidermal and mucosal bacterial
infections (Drake et al. 2008, Law et al. 1995b).
P. gingivalis, one of more than 600 bacterial species found in the oral cavity, is
among the most influential periodontal pathogens. This gram-negative, black-pigmented,
strict anaerobic coccobacillus is more likely to be found in patients with periodontitis and
less likely to be present in healthy individuals. Furthermore, P. gingivalis shows a strong
positive relationship with two parameters important in the diagnosis of periodontitis:
increased sulcular pocket depth and bleeding upon probing (Hutter et al. 2003, Socransky
and Haffajee 1992, Socransky et al. 1998). P. gingivalis is recognized as a late colonizer
in the development of oral biofilms (Kolenbrander et al. 2002, Socransky et al. 1998),
where the multitude of virulence factors produced by P. gingivalis contributes to its
pathogenicity (Holt et al. 1999). Additionally, P. gingivalis produces many proteins,
80
enzymes, and metabolic end products that are important to its survival and growth within
the host because they are active against a broad spectrum of host proteins and provide
mechanisms for evasion of host defenses (Holt et al. 1999).
Fatty acids (sapienic acid and lauric acid) and sphingoid bases (sphingosine and
dihydrosphingosine) present in the saliva and on the oral mucosa are particularly potent
against P. gingivalis with antimicrobial concentrations ranging from 0.2 – 125.0 µg/ml
(Table 4.1), and killing P. gingivalis cells in as little as six minutes (Figure 4.1).
Phytosphingosine, present in ceramides and glycosylceramides of oral epithelium, has
similar antimicrobial activity.
Within physiologic concentrations (4.0 - 13.2 µg/ml for total fatty acids and 0.5 –
5.0 µg/ml for sphingoid bases (Brasser et al. 2011a, Brasser et al. 2011b)), salivary lipids
exhibit potent antimicrobial activity against a variety of oral bacteria without apparent
harm to eukaryotic cells of the oral mucosa. However our lab recently showed that while
sphingoid bases are nontoxic to DCs at low concentrations they are extremely toxic to
DCs at higher concentrations (Figure 5.1; data and figure provided by Leslie Mahalick).
These results are similar to other studies showing dose-dependent cytotoxicity of
sphingosine and other sphingoid base derivatives against immortalized human endothelial
cells (HUVECtert cells) (Rozema et al. 2012), human epidermoid carcinoma KB cells
(Shirahama et al. 1997), murine P388 myeloid leukemia cells (Klostergaard et al. 1998),
and murine B16 melanoma cells (Rives et al. 2011). However, little is known about the
effects of sphingoid bases on normal (e.g. non-cancerous) human cells at cytotoxic versus
non-cytotoxic concentrations. We are interested in further exploring the effects of
sphingoid bases against DCs, particularly in the range of antimicrobial activity against P.
gingivalis that is within normal physiologic concentrations (Table 5.1).
81
Materials and Methods
Preparation of lipids
Sphingosine, dihydrosphingosine, and phytosphingosine were obtained from
Sigma Chemical Company (St Louis, MO) and glycerol monolaurate was obtained from
LKT Laboratories (St. Paul, MN). Lipids were prepared as described in Chapter 2 and
1.0 mg/ml stock solutions were diluted to the desired concentration using 0.14 M NaCl.
Glycerol monolaurate was used as a negative control because it is non-toxic to human
vaginal epithelial cells (Peterson and Schlievert 2006) and has not demonstrated
cytotoxicity for DCs in our studies even at high (80 µM) concentrations
Preparation of dendritic cells
Immature DCs were either obtained from AllCells (Emeryville, CA; PB-DC001F)
or prepared by treatment of human peripheral blood mononuclear cells (HPBMC; Lonza,
CC-2701; Walkersville, MD) with IL-4 (Peprotech, Rocky Hill, NJ) and granulocytemacrophage colony stimulating factor (GM-CSF; Peprotech, Rocky Hill, NJ) as
previously described (Sallusto and Lanzavecchia 1994). Monocytes were thawed,
suspended in LGM-3 (lymphocyte growth medium 3; Lonza; Walkersville, MD)
containing 1,000 U/ml IL-4 and 1,000 U/ml GM-CSF, placed in polysterene culture
flasks (Fisher Scientific, Pittsburg, PA), and incubated at 37ºC in an atmosphere
containing 5% CO2. After 50 minutes the media and non-adherent cells were replaced
with fresh LGM-3 with 1,000 U/ml IL-4 and 1,000 U/ml GM-CSF. Cells were incubated
for seven days at 5% CO2, adding fresh LGM-3 with 1,000 U/ml IL-4 and 1,000 U/ml
GM-CSF every other day. After one week, the monolayer of immature DCs was gently
washed from the flask with fresh LGM-3 containing 1,000 U/ml IL-4 and 1,000 U/ml
GM-CSF and centrifuged at 1,410 RPM (Eppendorf 5810 R; Hauppauge, NY) for ten
minutes to collect the floating and loosely adherent cells. Cells were tested for viability
using propidium iodide and resuspended in LGM-3 with 1,000 U/ml IL-4 and 1,000 U/ml
82
GM-CSF at 1 × 105 viable cells/ml; percentages of viable cells were recorded for each
experiment. A single cell culture was divided for flow cytometric and confocal
microscopic analyses of DCs with each sphingoid base treatment. Positive identification
of immature DCs was completed using four fluorochrome-conjugated antibodies for
CD1c, CD141, CD16, and CD11c (R&D Systems kit; Minneapolis, MN; data not shown)
(Piccioli et al. 2007). Killed DCs for use as a control were prepared by immersing tube
in 56° C waterbath for 10 minutes or by the addition of 2 mM hydrogen peroxide or 0.1%
sodium azide to the media.
Flow cytometry
For preparation of DCs for flow cytometry, a single cell culture was divided
among the appropriate number of tubes. A LIVE/DEAD® Cell Vitality Assay Kit
(L34951, Molecular Probes, Eugene, OR) utilizing C12-resazurin and SYTOX® Green
stains was used to differentiate between live and dead DCs after treatment with either
sphingosine, dihydrosphingosine, phytosphingosine, or glycerol monolaurate at
concentrations of 5 and 80 µM. This kit was chosen because it was reported to
differentiate between live, damaged, or dead cells based on the intensity of C12-resorufin
and SYTOX Green fluorescence in each cell. SYTOX Green is a fluorescent nucleic acid
stain that is impermeant to intact plasma membranes of live cells. C12-resazurin is
reported to be reduced to red-fluorescent C12-resorufin only in metabolically active cells
(Molecular Probes); however, preliminary experiments showed that C12-resazurin was
non-specifically taken up and reduced to C12-resorufin by all DCs regardless of their
dead/live status. Consequently, we were able to add both dyes prior to treatment, at
concentrations of 500 nM C12-resazurin and 10 nM SYTOX Green.
After the addition of dyes, sphingoid base treatments were added and an LSR II
Flow Cytometer (BD Biosciences, San Jose, CA) was used to measure the fluorescence
emission at times of 0, 20, 40, and 60 minutes, exciting at 488 (SYTOX Green) and 561
83
nm (C12-resorufin), and measuring emission at 530 and 585 nm. Compensation controls
were utilized to adjust for spectral overlap across dyes. Additional controls included
sphingoid bases in media with and without added dyes, untreated DCs, killed DCs, and
glycerol monolaurate. Flow cytometric data was analyzed using FlowJo software (Tree
Star, Inc, Ashland, OR) and all experiments completed in triplicate.
Confocal microscopy
For confocal microscopy, 600 µl of DCs resuspended in LGM-3 with 1,000 U/ml
IL-4 and 1,000 U/ml GM-CSF at 1 × 105 viable cells/ml were put into each of four
chambers of Lab-TekTM chamber slides (Thermo Scientific, Rockford, IL) and incubated
at 37ºC in an atmosphere containing 5% CO2 for two hours to allow attachment of cells.
The LIVE/DEAD stains described above in the flow cytometry section were added prior
to treatment at a concentration of 500 nM C12-resazurin and 10 nM SYTOX Green. After
addition of sphingoid base treatments, chamber slides were incubated for appropriate
times (0, 20, 40, and 60 minutes) before examination using a Zeiss 710 LSM (laser
scanning microscope) confocal microscope (Carl Zeiss Micro Imaging GmbH, Jena,
Germany). Controls included sphingoid bases in media with added dyes, glycerol
monolaurate-treated DCs, and untreated DCs.
Results
High concentrations (80 µM) of all the sphingoid bases quickly induced cell
damage leading to death of the DCs. While untreated DCs (Figure 5.2) lacked SYTOX
Green staining of their nuclei, exhibited normal healthy morphology with dendritic
processes extended, and remained attached to the slide, most sphingosine-treated DCs
immediately began to withdraw their dendritic processes (Figure 5.3A-B). Within 20
minutes (Figure 5.3C-D) many cells resembled the killed controls (Figure 5.4) with
SYTOX Green-stained nuclei, indicating compromise of the plasma membrane, and
detachment from the microscope slide. At 40 minutes (Figure 5.3E-F) and 60 minutes
84
(Figure 5.3G-H) incubation time, few cells remained attached to the slide and most of the
remaining cells’ nuclei were stained green. In addition, many cells appeared leaky and
distorted with cellular debris lying adjacent to them. Unexpectedly, C12-resazurin was
non-specifically taken up and reduced to red-fluorescent C12- resorufin by both dead and
live cells. Similar results were seen with 80 µM dihydrosphingosine treatment (Figure
5.5).
Cells treated with phytosphingosine (Figure 5.6) immediately exhibited
completely different morphologies depending upon where the sphingoid base pooled on
the slide. Cells lying adjacent to, or within a pool of phytosphingosine were immediately
killed before they could withdraw their dendritic processes (Figure 5.6B) while cells
further away from the pool of phytosphingosine (Figure 5.6A) died more slowly and
resembled cells treated with 80 µM treatments of either sphingosine or
dihydrosphingosine. These cells immediately began to withdraw their dendritic
processes and detach from the slide but were not immediately killed. Within 20 minutes
(Figure 5.6C-D) most cells had SYTOX Green-stained nuclei and at 40 minutes (Figure
5.6E-F) all cells were dead with some leaking of cellular contents (Figure 5.6E). Few
cells remained attached to the slide 60 minutes after phytosphingosine treatment and
those cells remaining were deteriorating, as evidenced by leaky, non-continuous plasma
membranes and breakdown of the nucleus (Figure 5.6H).
Treatment with 80 µM concentrations of the control lipid, glycerol monolaurate,
caused most of the cells to curl up immediately (Figure 5.7A-B). After 60 minutes
incubation most cells remained alive and attached to the slide, although they were still
rounded after incubation for one hour of incubation (Figure 5.7C-D).
All three sphingoid bases exhibited an affinity for the C12-resazurin dye (Figure
5.8) allowing easy visualization of lipid precipitate within the media. All of the
sphingoid bases in solution pooled in small drops scattered around the slide with some
precipitate within the lipid pools. Sphingosine precipitate (Figure 5.8A-B) appeared
85
stringy and aggregated in large clumps while phytosphingosine (Figure 5.8C-D), which
appeared as a crystalline precipitate, was suspended in much smaller aggregates.
Dihydrosphingosine (Figure 5.8E-F) was a flocculent precipitant. Glycerol monolaurate
appeared as varying sizes of lipid droplets without any precipitate and demonstrated little
affinity for C12-resazurin (Figure 5.8G-I). None of the lipids showed any affinity for the
SYTOX Green dye.
Sphingoid base affinity for C12-resazurin was also demonstrated using flow
cytometry (Figure 5.9) which highlighted the side/forward scatter pattern of each lipid
alone in media. This affinity for C12-resazurin was differential for each sphingoid base,
with sphingosine exhibiting the lowest fluorescence intensity and phytosphingosine
exhibiting an extremely high red fluorescent background that interfered with data
analysis.
Flow cytometry experiments also demonstrated the cytotoxic potential of
sphingoid bases. All three sphingoid bases were cytotoxic to DCs at 80 µM
concentrations (Figure 5.10), quickly killing a large percentage of cells. Within 20 – 40
minutes post-treatment with sphingosine and dihydrosphingosine flow cytometric data
showed that the majority of DCs were dead (Figure 5.11) while treatment with 5 µM
sphingoid base concentrations were not cytotoxic (e.g. sphingosine) or only mildly
cytotoxic (e.g. dihydrosphingosine and phytosphingosine) for DCs. The control lipid,
glycerol monolaurate, was not cytotoxic at any of the concentrations tested. Surprisingly,
despite a shift in flow cytometry side/forward scatter graphs (Figure 5.12) usually
indicative of cell death, the percentage of dead cells with phytosphingosine and
dihydrosphingosine (data not shown) treatment indicated by flow cytometry (Figure 5.11)
was not consistent with 100% DC death.
Examination of DCs by confocal microscopy also demonstrated the low
cytotoxicity of sphingoid bases at lower concentrations (5 µM). Upon treatment with
sphingosine (Figure 5.13), dihydrosphingosine (Figure 5.14), or phytosphingosine
86
(Figure 5.15), most cells were still alive one hour after treatment although many cells
reacted to low lipid concentrations by withdrawing their dendritic processes. Although
treatment of DCs with 5 µM concentrations of dihydrosphingosine and phytosphingosine
caused DCs to curl up and caused a few cells to detach, most cells remained alive one
hour after treatment. After treatment with 5 µM concentrations of glycerol monolaurate
(Figure 5.16) DCs remained unaffected and were similar to untreated controls (Figure
5.2).
Discussion
With the discovery of sphingoid base antimicrobial activity towards P. gingivalis,
it is important to establish the cytoxocity of sphingoid bases against host cells,
particularly in the range of antimicrobial activity against P. gingivalis that falls within
normal physiologic concentrations (e.g. 5-20 µM; (Brasser et al. 2011a, Brasser et al.
2011b). For this experiment, we chose DCs to begin our study of the cytotoxicity of
sphingoid bases against host cells because DCs are the primary immune cell that would
come into contact with lipids in the epithelium. Although 5 µM concentrations of
sphingosine, dihydrosphingosine, and phytosphingosine do elicit a response (e.g. DCs
withdraw their dendritic processes upon exposure), they are not cytotoxic to DCs within
an hour of treatment and do not induce cellular damage, as indicated by the lack of
SYTOX Green uptake. This is supported by previous experiments in our lab in which 5
µM concentrations of sphingosine, dihydrosphingosine, and phytosphingosine were not
cytotoxic to DCs within 24 hours after treatment.
DCs are much more susceptible to high sphingoid base concentrations than we
expected. In this study 80 µM concentrations of sphingosine, phytosphingosine, and
dihydrosphingosine, but not glycerol monolaurate, were cytotoxic to DCs, inducing
cellular damage and death in less than 20 minutes for most treatments. These results are
similar to previous studies in our lab showing that DCs treated with 40 – 80 µM
87
concentrations of sphingosine, dihydrosphingosine, or phytosphingosine are cytotoxic to
normal human DCs within 24 hours while glycerol monolaurate is non-toxic to DCs even
at 80 µM concentrations (work completed by Leslie Mehalick). Sphingoid bases are
cytotoxic for other eukaryotic cell types as well and several sphingoid base derivatives
have been selected for their cytotoxic properties and studied as potential treatments for
cancer cells (Klostergaard et al. 1998, Rives et al. 2011, Rozema et al. 2012, Shirahama
et al. 1997).
Because C12-resazurin was non-specifically taken up and reduced to C12-resorufin
whether cells were dead or alive, it was not a good indicator of cellular metabolism;
therefore we excluded it from flow cytometry analyses and focused on the uptake of
SYTOX Green dye. However, the affinity of all three sphingoid bases for C12-resazurin
still created some difficulty in this study. Lipid-treated samples had a very high red
fluorescent background due to excess C12-resazurin-dyed lipid in the samples – especially
with high lipid concentrations of phytosphingosine and dihydrosphingosine. Although a
wash of the cells would have removed excess lipids from the media, this was not possible
given the longitudinal design of our experiments. Because the scatter pattern of these
dyed lipids was similar to that of dead DCs, it was difficult to gate out the dyed sphingoid
bases without also excluding some of the dead cells. Furthermore, because DCs quickly
take up lipids (Shirahama et al. 1997) the red fluorescence intensity of the DCs after
treatment with C12-resazurin-stained sphingoid bases was greatly increased, which
created further problems both with confocal imaging and with flow cytometry. The main
drawback of using SYTOX Green as a dead/live indicator in confocal microscopy is that
images must be taken very quickly due to rapid photobleaching.
When examined by confocal microscopy, most cells treated with 80 µM
concentrations of all three sphingoid bases were dead within 40 minutes. Flow
cytometric analysis of DCs treated with 80 µM dihydrosphingosine and
phytosphingosine, however, did not support the expected percentage of dead cells,
88
although a visible shift in the forward\side scatter plot indicated that the majority of the
cells were dead or dying. Several potential explanations were explored. Gating error
could contribute to an error in calculation of dead and live cell percentages; it is possible
that our attempt to gate out dyed excess lipids resulted in gating out too many dead cells.
However, when we did not gate out the dyed lipids, the percentage of dead cells was still
much lower than expected. It is also possible that dead DCs were not taking up SYTOX
Green, but our confocal images do not support this theory. Another possible explanation
is that the cells treated with high concentrations of sphingoid bases undergo a slow lytic
process and therefore a percentage of them are not present for flow cytometric analysis.
This theory is supported by several observations including increased amounts of debris
that were gated out of the analyses and by confocal evidence of cell deterioration seen,
especially with dihydrosphingosine and phytosphingosine treatments. In addition,
previous data from our lab (completed by Leslie Mehalick) shows an increase in the
release of lactate dehydrogenase (LDH) upon treatment with sphingoid bases that is not
present in cells killed with sodium azide, indicating cellular damage and/or cell lysis.
Finally, with all 80 µM sphingoid base treatments, detachment of cells from the
microscope slides was evident but unattached cells were not plentiful on the slide. It is
therefore possible that treated DCs detach and slowly lyse, leading to their death and
decreased detection by flow cytometry.
The concentration-dependent effect of sphingoid bases against both host cells and
bacteria highlights the significance of balanced and controlled lipid concentrations in
saliva. Lipid deficiencies or imbalances in lipid ratios are associated with several
diseases. For example, both deficient hexadecanoic acid production (Takigawa et al.
2005) and decreased levels of sphingosine (Arikawa et al. 2002) are associated with
atopic dermatitis and subsequent increase in S. aureus skin colonization within otherwise
healthy individuals. Furthermore, cystic fibrosis is linked with deficient fatty acid
production (Freedman et al. 2004, Strandvik et al. 2001). In another study, mutation of
89
an enzyme necessary for palmitic and oleic acid production was linked to failure to clear
skin infections of S. aureus or Streptococcus pyogenes within innate immunodeficient
mice (Georgel et al. 2005). It is therefore possible that imbalances in lipid ratios or
defective production of certain lipids could be responsible for increased ability of P.
gingivalis to colonize the oral cavity, potentially leading to increased periodontal disease.
Interestingly, although these physiologic concentrations of sphingoid bases tested here
are not cytotoxic to DCs, they are antimicrobial to a variety of both gram-positive and
gram-negative bacteria (Bergsson et al. 2001a, Bergsson et al. 2002, Bibel et al. 1992b,
Bibel et al. 1993, Burtenshaw 1942, Fischer et al. 2012, Fischer et al. 2013) including P.
gingivalis.
It becomes reasonable then to speculate that topical application of endogenous
lipid formulations could potentially supplement the natural immune function of lipids on
mucosal surfaces and in saliva, restoring a healthy lipid balance to reduce or treat
infections (Thormar and Hilmarsson 2007). Drake, et al. (Drake et al. 2008) point out
that because they are normal occupants of the skin [and oral mucosa] these lipids are
likely to be less irritating. In addition, because of their evolution with the potential
pathogens of skin and oral mucosa it is more unlikely that these pathogens will readily
develop resistance to treatment (Drake et al. 2008). Crucial to the development of
formulations that would stimulate the natural innate function is a better understanding of
the spectrum of sphingoid base cytotoxic effects on eukaryotic cells. This study
contributes to that knowledge base and indicates a range of sphingoid base concentrations
that would be effective antimicrobial agents against bacteria without harm to host cells.
90
µM Sphingosine Phytosphingosine Dihydrosphingosine
µg/ml
µg/ml
µg/ml
160
48.1
50.8
48.3
80
24.1
25.4
24.2
40
12.0
12.7
12.1
20
6.0
6.4
6.0
10
3.0
3.2
3.0
5
1.5
1.6
1.5
2.5
0.8
0.8
0.8
1.3
0.4
0.4
0.4
0.6
0.2
0.2
0.2
0.3
0.1
0.1
0.1
Cytotoxic for DC
Antimicrobial for
P. gingivalis
Table 5.1. Antimicrobial and cytotoxic activity of sphingoid bases.
Note: Shown here is the the overlap of sphingoid base concentrations that are
antimicrobial for P. gingivalis (highlighted in yellow) and non-cytotoxic for DCs
within the range of physiologic concentrations (green digits). Cytotoxic
concentrations sphingoid bases are outlined in red.
91
Figure 5.1. Cytotoxicity of sphingoid bases against DCs. Cytoxicity of sphingoid
bases against DCs was measured using three separate assays: 1) alamar blue,
which detects metabolic activity; 2) propidium iodidide (PI), the uptake of
which indicates a damaged cellular membrane; and 3) the release of lactate
dehydrogenase (LDH) from dead/damaged cells. Controls include glycerol
monolaurate, as a non-cytotoxic lipid control and sodium azide-killed DCs
(KC). MFI = mean fluorescence intensity. LC = live, untreated DCs.
Note: Data and figure used with permission from Dr Leslie Mehalick.
92
Figure 5.2. Confocal micrographs of untreated DCs. Untreated DCs indicated mostly
live, healthy cells, indicated by C12-resazurin staining and lack of SYTOX
Green-stained nuclei. C12-resazurin is reported to be reduced to redfluorescent C12-resorufin only in metabolically active cells (Molecular Probes)
while SYTOX Green is a green-fluorescent nucleic acid stain that is
impermeant to intact plasma membranes of live cells. Scale bars = 10 µm.
Figure 5.3. Confocal micrographs of DCs treated with 80 µM sphingosine. DCs treated with 80 µM sphingosine and visualized
immediately (A, B) have started to curl up and detach from the slide. By 20 minutes (C, D) many cells exhibit damaged
plasma membranes, as evidenced by the ability of SYTOX Green to permeate the cell and stain the nuclei. At 40 (E, F)
and 60 minutes (G, H) most cells are dead and detaching, leaving few cells attached to the slide. Scale bars = 10 µm.
93
94
Figure 5.4. Confocal micrographs of killed DCs. DCs were heat-killed (56° C
waterbath for 10 minutes) or killed with either 2 mM hydrogen peroxide or
0.1% sodium azide followed by the addition of C12-resazurin and SYTOX
Green fluorescent dyes as described above. All cells were dead, as evidenced
by SYTOX Green-stained nuclei, indicating compromise of the plasma
membrane. Although C12-resazurin is reported to be reduced to redfluorescent C12-resorufin only in metabolically active cells these experiments
showed that C12-resazurin was non-specifically taken up and reduced to C12resorufin by all DCs regardless of their live/dead status. Scale bars = 10 µm.
95
Figure 5.5. Confocal micrographs of DCs treated with 80 µM dihydrosphingosine. DCs treated with 80 µM dihydrosphingosine
and visualized immediately (A, B) and at 20 minutes (C, D) have started to curl up and detach from the slide. Within 40
minutes (E, F) many cells exhibit damaged plasma membranes, as evidenced by the ability of SYTOX Green to permeate
the cell and stain the nuclei. At 60 minutes (G, H) most cells are dead and detaching, leaving few cells on the slide. Scale
bars = 10 µm.
96
Figure 5.6. Confocal micrographs of DCs treated with 80 µM phytosphingosine. DCs treated with 80 µM phytosphingosine and
visualized immediately (A,B) exhibited two different morphologies depending upon the location of the cells relative to the
“pooling” of phytosphingosine on the slide. Cells with phytosphingosine pooled around them were dead immediately after
treatment (B) while cells without lipid pooled directly around them (A) immediately started rounding up and detaching
from the slide. Within 20 minutes (C, D) most cells exhibit SYTOX Green-stained nuclei and phytosphingosine can be
visualized adjacent to some cells (D, white arrow). At 40 (E, F) and 60 minutes (G, H) most cells are dead, deteriorating,
and detaching, leaving few cells attached to the slide.
97
Figure 5.7. Confocal micrographs of DCs treated with 80 µM glycerol monolaurate.
DCs treated with an 80 µM concentration of glycerol monolaurate and
visualized immediately (A, B) have withdrawn their dendritic processes but
stay attached to the slide and even at 60 minutes (C, D) do not exhibit SYTOX
Green-stained nuclei. Scale bars = 10 µM.
98
Figure 5.8. Confocal micrographs showing sphingoid base affinity for C12-resazurin dye. Sphingosine (A, B), phytosphingosine
(C, D), and dihydrosphingosine (E, F) all showed an affinity for C12-resazurin dye. Glycerol monolaurate (G-I) showed
almost no affinity for the red fluorescent dye. None of the lipids appeared to take up any SYTOX Green dye. Glycerol
monolaurate is a non-cytotoxic lipid used as a control.
Figure 5.8. Continued
99
Dihydrosphingosine
Phytosphingosine
Glycerol Monolaurate
5 µM
80 µM
Sphingosine
Figure 5.9. Flow cytometry data showing sphingoid base affinity for C12-resazurin dye. Flow cytometric analyses of sphingoid
bases and glycerol monolaurate in media with added C12-resazurin and SYTOX Green dyes indicated that all three
sphingoid bases were differentially dyed by C12-resazurin as seen by the intensity of the scatter pattern in each figure. No
SYTOX Green fluorescence was detected for any of these lipids (data not shown). C12-resazurin-dyed phytosphingosine at
80 µM exhibited the highest intensity fluorescence, while glycerol monolaurate at 80 µM showed almost no affinity for the
red fluorescent dye.
100
20 min
40 min
60 min
Killed
Untreated
0 min
Figure 5.10. Flow cytometric analyses of sphingoid base cytotoxicity against DCs. Flow cytometric data for treatment of DCs
with sphingosine (Sph), dihydrosphingosine (Dih), phytosphingosine (Phy), and glycerol monolaurate (GM) at high (80
µM) and low (5 µM) concentrations shows that high concentrations of all three sphingoid bases are cytotoxic within 20
minutes as indicated by the shift of the SYTOX Green peak. Lower concentrations of these sphingoid bases are not
cytotoxic to DCs and glycerol monolaurate is not cytotoxic to DCs at any of the concentrations tested.
101
20 min
40 min
60 min
Dih (80 µM)
Sph (80 µM)
0 min
Figure 5.10. Continued
102
20 min
40 min
60 min
GM (80 µM)
Phy (80 µM)
0 min
Figure 5.10. Continued
103
20 min
40 min
60 min
Dih (5 µM)
Sph (5 µM)
0 min
Figure 5.10. Continued
104
0 min
20 min
40 min
60 min
GM (5 µM)
Phy (5 µM)
0
Figure 5.10. Continued
105
106
A
B
C
Figure 5.11. Cytotoxicity of sphingoid bases for DCs. Graphical representation of flow
cytometric data for the treatment of DCs with sphingosine (A),
dihydrosphingosine (B), phytosphingosine (C), and glycerol monolaurate (AC) at high (80 µM) and low (5 µM) concentrations shows cytotoxicity of each
lipid over a one-hour time period. Each time point represents an average of
three individual longitudinal experiments. Error bars represent SEM; where
no error bars are present the SEM was zero.
107
20 min
40 min
60 min
0
Phy (80 µM)
Killed Cells
Untreated
0 min
Figure 5.12. Forward/side scatter graphs of flow cytometric data for treatment of
DCs with phytosphingosine (Phy) at 80 µM concentrations shows the shift
typically indicating the death of cells.
108
Figure 5.13. Confocal micrographs of DCs treated with 5 µM sphingosine. Many DCs treated with a 5 µM concentration of
sphingosine immediately begin to curl up (A, B) and continue to withdraw their dendritic processes as time passes, seen at
20 (C, D), 40 (E, F), and 60 minute (G, H) incubation times. However, very few cells have damaged cellular membranes,
as evidenced by the lack of SYTOX Green nuclear staining. Scale bars = 10 µm.
109
Figure 5.14. Confocal micrographs of DCs treated with 5 µM dihydrosphingosine. Many DCs treated with a 5 µM concentration
of dihydrosphingosine immediately begin to curl up (A, B) and continue to withdraw their dendritic processes as time
passes, seen at 20 (C, D), 40 (E, F), and 60 minute (G, H) incubation times, but very few have damaged cellular
membranes, as evidenced by the lack of SYTOX Green nuclear staining. Scale bars = 10 µm.
110
Figure 5.15. Confocal micrographs of DCs treated with 5 µM phytosphingosine. DCs treated with a 5 µM concentration of
phytosphingosine and visualized immediately (A, B) have withdrawn their dendritic processes. As time passes this trend
can be seen at times of 20 (C, D), 40 (E, F), and 60 minutes (G, H), but very few exhibit damaged cellular membranes, as
evidenced by the lack of SYTOX Green nuclear staining. Scale bars = 20 µm.
111
Figure 5.16. Confocal micrographs of DCs treated with 5 µM glycerol monolaurate.
DCs treated with a 5 µM concentration of glycerol monolaurate and visualized
immediately (A, B) and at 60 minutes (C, D) appear similar to untreated
controls (Figure 5.2). Scale bars = 10 µm.
112
CHAPTER 6
CONCLUSIONS
In this study I have described the differential antibacterial effect of lipids
endogenous to the oral cavity against a variety of bacterial species present in the oral
cavity and on skin. The dose-dependent antibacterial activity of these sphingoid bases
and fatty acids is both bacteria-specific and lipid-specific. All of the treatment lipids are
quickly taken up in large quantities by each bacterial species tested, and cause visible
intracellular and extracellular damage. In P. gingivalis, sapienic acid induces changes in
the proteome. While fatty acids exhibited a broad range of antimicrobial activity that
varied from very potent (e.g. F. nucleatum) to mildly or non-inhibitory (e.g. Cornyeform
bacteria), sphingoid bases were very potently inhibitory against all the bacteria tested
except for P. aeruginosa and S. marcescens. Interestingly, fatty acids were more active
against oral bacteria than any other bacteria tested and sphingoid bases were most active
against P. gingivalis with MICs ranging from 0.2 to 0.8 µg/ml. Overall, the variability of
sphingoid base and fatty acid antimicrobial activity against all bacteria was highly
significant in most cases.
Potential Mechanisms of Activity
The dose-dependent and specific antimicrobial activity exhibited by each of these
oral lipids lends credence to the proposal that sphingoid bases and fatty acids serve an
innate immune function in the oral cavity. An extensive number of host innate immune
factors, including anionic peptides (Brogden et al. 1996), cathelicidins (Kalfa et al. 2001),
and defensins (Harder et al. 2001, Shimoda et al. 1995), induce extensive damage to
gram-positive and gram-negative bacteria similar to what I have described here. Activity
of these previously described innate immune factors depends upon the size of the
molecule, specific amino acid sequences, charge, structural conformation,
hydrophobicity, and amphipathicity (Brogden 2005) and mechanisms of action include
113
flocculation of intracellular contents, alteration of the bacterial cytoplasmic membrane
(e.g. pore formation), or inhibition of various cellular process (e.g. enzymatic activity and
cell wall, nucleic acid, or protein synthesis) (Brogden 2005).
Although discussion of specific mechanisms of action for sphingoid bases is
sparse, potential mechanisms of fatty acid antimicrobial activity have been reviewed and
are discussed in Chapter 4. These include creation of pores in the bacterial cell, alteration
of the cellular membrane, lysis of the cell, and disruption of various cellular processes
either by interference of spatial arrangement or by direct binding to proteins (Desbois and
Smith 2010). In this study I show evidence of cellular membrane disruption, bacterial
cell lysis, and flocculation of intracellular contents. In addition, these studies provide
evidence that sphingoid bases and fatty acids are taken up by all the bacteria tested.
Furthermore, treatment of P. gingivalis with sapienic acid induces upregulation of a set of
proteins involved in various cellular processes. Proteomic changes induced by different
stress situations have been well studied in P. gingivalis (See Appendix A) and each
stressor induces a unique response with very little overlap. I propose that treatment of P.
gingivalis sapienic acid induces a unique stress response in this bacterium, which is
evidenced by the upregulation of a unique set of proteins.
There are several potential mechanisms of activity for sphingoid bases and fatty
acids against P. gingivalis. In addition, lipid structure differences (discussed in Chapter
4) potentially affect mechanism; therefore, mechanisms of action are likely complicated
and could include a different mechanism for each lipid. Based on demonstrated
differential activity of sphingoid bases and fatty acids against various bacteria, it is likely
not a basic surfactant effect. Lipids may be transported into the cell and accumulate in
the cytoplasm where they can interact with various protein components of the cytosol,
inhibiting cytosolic enzymes or bacterial fatty acid synthesis (Desbois and Smith 2010).
On the other hand, lipids may insert into the membranes of bacteria causing changes in
the physical properties of the membrane (e.g. membrane fluidity, size, shape) which
114
could potentially disrupt energy production through spatial orientation of electron
transport chain components (Desbois and Smith 2010).
The combined data presented in this study suggest that with sapienic acid, there
may be a quick two-step process leading to antimicrobial activity that appears to be time
and sapienic acid concentration dependent. As P. gingivalis cells are exposed to sapienic
acid they start taking up large amounts of the lipid, become stressed and quickly mount a
response by adjusting protein activity, as evidenced by the differential protein profiles
and the upregulation of several components important in microbial metabolism in diverse
environments. It is possible, however, that as a critical point (time and/or lipid
concentration) is reached, rescue attempts fail as these cells succumb to lysis. Further
analysis of the metabolic consequences of sapienic acid treatment on P. gingivalis will be
necessary to confirm this and could be the subject of future studies.
Implications for Prophylactic or Therapeutic Treatments
With the expanding resistance of bacteria to many available antibiotic treatments
(Thormar and Hilmarsson 2007) alternative treatments are becoming increasingly
important. In addition, it is also important to demonstrate that these compounds could be
used without harm to the host. Sphingoid bases are cytotoxic to many types of eukaryotic
cells, however they are not toxic to DCs at physiologic concentrations nor at
concentrations which are antimicrobial to the bacterial species we tested. Therefore, they
appear to be safe and efficacious at their antimicrobial does and could therefore have
potential for prophylactic or therapeutic intervention of infection.
115
APPENDIX
Gene
Acc #
hprA
Protein/Function
Heat Oxidative
(O2)
Oxidative
(H2O2)
pH
Heme
limitation
EtOH
Epithelial
contact
Glycerate
dehydrogenase
^
SPg1
PG0184
Transposase
^
˅
^
nfB
PG0255
Translation initiation
factor IF-2
^
˅
^
gpA
rtT
PG2024
PGN_
1970
Arg-specific cysteine
proteinase
^˅
^
˅
^
gpB
PG0506
Arg-specific cysteine
proteinase
^
^ ˅
^
^
^
^ ˅
^
^
PGN_
1466
kgp
Kgp; Lys-gingipain
(cysteine protease)
Sapienic
acid
˅
^
^
^
Table A.1. P. gingivalis stress responses.
116
Note: Literature review of P. gingivalis upregulated and downregulated proteins in stress responses associated with heat stress
(Amano et al. 1994, Bonass et al. 2000, Lopatin et al. 1999, Lu and McBride 1994, Murakami et al. 2004, Percival et al. 1999,
Shelburne et al. 2005, Vayssier et al. 1994), O2 oxidative stress (Meuric et al. 2008, Shelburne et al. 2005, Vayssier et al. 1994),
H2O2 oxidative stress (Meuric et al. 2008, Shelburne et al. 2005, Vanterpool et al. 2010), pH stress (Lu and McBride 1994,
Vayssier et al. 1994), heme limitation (Dashper et al. 2009), EtOH stress (Lu and McBride 1994), and response to contact with
epithelial cells (Hosogi and Duncan 2005). Upregulation is indicated by ^, while downregulation is indicated by ˅.
Gene
Acc #
Protein/Function
Heat Oxidative
(O2)
yrG
PG0525
CTP synthase
^
roEL
PG0520
HSP60
^
mpF
PG0695
Immunoreactive
antigen P32, porin
^
˅
^
YP2
PG0774
Hypothetical protein
^
˅
^
psC
PG1135
Glycosyltransferase
^
˅
^
rpsA
PG1297
Ribosomal protein S1
^
^
^
sodB
PG1545
Superoxide dismutase
Fe-Mn
^
^ ˅
^
humY
PG1551
HmuY protein
^
˅
^
secDF
PG1762
Protein-export
membrane protein
^
^ ˅
^
HYP1
PG1795
43-64 kDa LPSmodified surface
protein (P27)
^
^ ˅
^
cysS
PG1878
Cysteinyl-tRNA
synthetase
^
˅
^
htpG
PG0045
naK
PG1208
Heat shock protein 70
abfD
PG0692
4-hydroxy-CoA
dehydratase
^
Heme
limitation
EtOH
Epithelial
contact
^
^
Sapienic
acid
^
^
^
^
^
^
˅
pH
^
^
^
^
^
˅
117
Table 1. Continued
˅
Oxidative
(H2O2)
Gene
Acc #
fimA
Protein/Function
Heat Oxidative
(O2)
pH
Heme
limitation
Sapienic
acid
^
PG0618
Alkyl hydroperoxide
reductase C
^
phF
PG0619
Alkyl hydroperoxide
reductase F
^
ps
PG0090
Dps family protein
(DNA-binding protein
from starved cells)
^
atA
PG1582
BatA protein
^
tn
PG1286
Ferritin
^
px
PG1729
Thiol peroxidase
^
˅
^
^
˅
˅
^
^
Hemagglutinin-like
protein
^
PG0548
Pyruvate
ferredoxin/flavodoxin
oxidoreductse family
protein
^
PG0687
Succinatesemialdehyde
dehydrogenase, AFD
^
118
Table 1. Continued
Epithelial
contact
^
aphC
sucD
EtOH
˅
FimA, structural
subunit of fimbriae
HSP90 homologue
Oxidative
(H2O2)
Gene
Acc #
Protein/Function
hbD
PG0689
NAD-dependent 4hydroxybutyrate
dehydrogenase
^
PG0690
4-hydroxybutyrateCoA transferase
^
PG0691
NifU-like protein
^
PG0692
4-hydroxybutyryl-CoA
dehydratase
^
PG1067
Hypothetical protein
^
PG1068
Conserved
hypothetical protein
^
acdA
PG1076
Acyl-CoA
dehydrogenase, shortchain specific
^
ackA
PG1081
Acetate Kinase
^
pta
PG1082
Phosphotransacetylase
^
gdh
PG1232
Glutamate
dehydrogenase, NADspecific
abfD
Heat Oxidative
(O2)
^
Oxidative
(H2O2)
pH
Heme
limitation
^
EtOH
Epithelial
contact
Sapienic
acid
^
Table 1. Continued
119
Gene
Acc #
Protein/Function
frdB
PG1614
Fumarate reductase,
iron-sulfur protein
(FrdB)
˅
frdA
PG1615
Fumarate reductase,
flavoprotein subunit
(FrdA)
˅
PG0026
Hypothetical protein
(homology to Arg
proteases)
^
PG0159
Endopeptidase PepO
^
PG0350
Internalin-related
protein
^
PG1374
Immunoreactive 47kDa protein (IrpI)
^
PG2130
FimX
^
PG2131
PgmA
^
PG2132
Fimbrillin FimA
^
PG2133
Lipoprotein
^
PG2134
FimC
^
PG2135
FimD
^
PG2136
FimE
^
pepO
fimA
Oxidative
(H2O2)
pH
Heme
limitation
EtOH
Epithelial
contact
Sapienic
acid
˅
˅
120
Table 1. Continued
Heat Oxidative
(O2)
Gene
Acc #
Protein/Function
PG1638
Thioredoxin family
protein
^
PG1639
Hypothetical protein
^
PG1640
DinF, membranespanning MATE
efflux pump
^
PG1641
PtpA, protein tyrosine
phosphatase
^
PG1642
Cation-translocating
ATPase (ZntA)
^
PG0616
Thioredoxin/HBP35
(heme-binding
protein)
^
PG0618
Alkyl hydroperoxide
reductase
^
PG0619
Alkyl hydroperoxide
reductase subunit F
^
PG0644
HtrE (Tla); TonBlinked receptor
^
PG0645
HtrD; no known
function
^
PG1043
FeoB2
^
Oxidative
(H2O2)
pH
Heme
limitation
EtOH
Epithelial
contact
Sapienic
acid
121
Table 1. Continued
Heat Oxidative
(O2)
Gene
hmuR
Acc #
Protein/Function
PG0646
HtrC; ABC heme
transport system ATP
binding protein
^
PG0647
HtrB; ABC heme
transport system
permease
^
PG0648
HtrA; ABC heme
transport system solute
binding protein
^
PG1551
HmuY
^
PG1552
HmuR
^
PG1553
HmuS
^
PG1554
Hypothetical protein
^
PG1555
TolQ
^
Pg1556
HmuV
^
PG1019
Hypothetical protein
^
PG1020
Conserved
hypothetical protein;
possible outer
membrane receptor
protein
^
PG1858
Flavodoxin A
^
Oxidative
(H2O2)
pH
Heme
limitation
EtOH
Epithelial
contact
Sapienic
acid
122
Table 1. Continued
Heat Oxidative
(O2)
Gene
Acc #
Protein/Function
Heat Oxidative
(O2)
PG1874
Conserved
hypothetical protein
^
PG1875
Hemolysin A
^
31 and 26 kDa
proteins (putative
function: heme
uptake/storage)
60 and 68 kDa
proteins
pH
Heme
limitation
EtOH
Epithelial
contact
^
^
^
htpG
PG0045
Heat shock protein 90
^
htrA
PG0593
Heat-induced serine
protease
^
slyD
PG1315
FKBP-type
peptidylprolyl
isomerase
^
grpE
PG1775
Heat shock protein
^
dnaJ
PG1776
Heat shock protein 40
^
trxB
PG1134
Thioredoxin reductase
^
123
Table 1. Continued
Sapienic
acid
˅
19 and 50 kDa
proteins
92 and 80 kDa
proteins
Oxidative
(H2O2)
Gene
Acc #
Protein/Function
cpP
PG1765
Acyl carrier protein
^
fabG
PG1239
Acyl carrier protein
^
abF
PG1764
3-Oxoacyl synthase
^
abH
Heat Oxidative
(O2)
Oxidative
(H2O2)
pH
Heme
limitation
EtOH
Epithelial
contact
3-oxoacyl-[acyl carrier
protein] synthase 3
^
^
prY
PG1089
Transcriptional
regulator
^
etR
PG1240
Transcriptional
regulator
^
mpH1
PG0192
Cationic outer
membrane protein
^
mpH1
PG0193
Cationic outer
membrane protein
^
PG0435
Capsule biosynthesis
^
PG2167
Immunoreactive 53kDa antigen
^
PG0419
Conserved
hypothetical
^
PG0686
Conserved
hypothetical
^
PG0434
Hypothetical
^
124
Table 1 Continued
Sapienic
acid
Gene
groES
Acc #
Protein/Function
Heat Oxidative
(O2)
Oxidative
(H2O2)
PG0654
Hypothetical
^
PG1316
Hypothetical
^
PG1317
Hypothetical
^
PG1635
Hypothetical
^
PG0521
pH
Heme
limitation
EtOH
Epithelial
contact
Sapienic
acid
^
ragA
RagA (Pgm1); 110
kDa
˅
ragB
RagB (Pgm4); 47-55
kDa
˅
75 kDa protein (Pgm2)
˅
Fimbrial proteins
˅
Arg-X
serC
phosphoserine
aminotransferase
gapA
Glyceraldehyde 3phosphate
dehydrogenase, type 1
^
^
^
^
^
^
Table 1. Continued
125
126
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