Impacts of Mucins on Microbial Physiology and
Interactions
by
Nicole Lynn Kavanaugh
B.S. Biology
Hofstra University, 2008
Submitted to the Microbiology Graduate Program in partial fulfillment of the requirements for
the degree of Doctor of Philosophy at the Massachusetts Institute of Technology
June 2015
© 2015 Nicole Kavanaugh. All Rights Reserved.
The author hereby grants to MIT permission to reproduce and to distribute publicly paper and
electronic copies of this thesis document in whole or in part in any medium now known or
hereafter created.
Author ……………………………………………………………………………………………...
Nicole L. Kavanaugh
Microbiology Graduate Program
May 22, 2015
Certified by………………………………………………………………………………………...
Katharina Ribbeck
Assistant Professor of Biological Engineering
Thesis Supervisor
Accepted by………………………………………………………………………………………...
Michael Laub
Professor of Biology
Director Microbiology Graduate Program
2
Impacts of Mucins on Microbial Physiology and
Interactions
by
Nicole Lynn Kavanaugh
Submitted to the Microbiology Graduate Program in partial fulfillment of the requirements for
the degree of Doctor of Philosophy at the Massachusetts Institute of Technology
ABSTRACT
The human body is colonized by trillions of microbes known collectively as the
microbiota. Many of these organisms inhabit mucosal surfaces, with most found in the large
intestines, but many also dwell in the respiratory and urogenital tracts. Despite the enormous
microbial population inhabiting the body, some of which are opportunistic pathogens, most
people harbor these organisms without showing any signs of disease. The mucus that covers the
wet epithelium and houses the microbiota is a prime candidate for offering protection from
pathogens, yet its specific role is poorly understood.
The object of this thesis is to explore the effects of mucins, the main gel-forming
component of mucus, on microbial behavior. Using an in vitro mucus model consisting of
purified mucins, I show that these polymers suppress microbial virulence traits in selected
bacterial and fungal opportunistic pathogens and influence the composition of microbial
communities.
In Chapter 2, I study the impact of mucins on bacterial biofilm formation using the
opportunistic pathogen Pseudomonas aeruginosa. I demonstrate that mucins reduce surface
attached biofilm formation. However, P. aeruginosa can overcome mucin-induced biofilm
suppression if flagellar motility is lost, allowing them to form non-surface attached biofilms that
are suspended in mucins.
In Chapter 3, I analyze the effects of mucins on the virulence traits of Candida albicans,
a fungal opportunistic pathogen. The results show that mucins broadly suppress virulence traits
of this organism, such as surface attachment, hyphal formation and biofilm formation, at both the
levels of gene expression and phenotype.
In Appendix A, I combine P. aeruginosa and C. albicans with mucins to determine the
influence of these molecules on interspecies interactions. Whereas P. aeruginosa typically kills
C. albicans, mucins protect the fungus from bacterial pathogenicity. Therefore, in addition to
influencing microbial virulence, mucins can impact microbial community dynamics.
Overall, my work suggests that mucins protect the body from microbes by functioning as
a modulator of microbial behavior, coercing certain microbes to downregulate virulence gene
and trait expression, thereby influencing microbial effects on the host and within the microbiota.
Thesis Supervisor: Katharina Ribbeck
Title: Assistant Professor of Biological Engineering
3
4
Acknowledgements
First and foremost I would like to thank my thesis mentor Katharina Ribbeck. The passion that
Katharina shows for her work is what inspired me to join her lab. She was always there when I
needed advice but also gave me generous freedom to pursue my ideas. For this, I will always be
grateful.
I would like to thank my past and present thesis committee members, Chris Kaiser,
Gerry Fink, Tim Lu and Jacquin Niles, as well as Katherine Lemon for joining from the Forsyth
Institute. I am very thankful for your time as well as your indispensible advice throughout the
years. Thank you to Clarissa Nobile, who has been a tremendously supportive collaborator. I am
also very grateful to Alan Grossman for his encouragement when I needed it most.
A very special thank you to all the Ribbeckers. Thank you for listening when I wanted to
run an idea by you, for always being willing to get coffee or to eat way too much food, and for
being great friends. Thank you to Thomas, Nicole B. (AKA “Tall Nicole”), Andrew, Tu, Julia,
Leon, Erica, Kate, Wes, Tahoura, and Brad. A special thank you to the original crew, Marina,
Oliver, Alex and Regi, an amazing group that helped me get acclimated when I first joined the
Ribbeck Lab. I would also like to thank the undergraduate researchers that worked with me,
Anna, Emily and Angela. A special thanks to Nicole B, Julia, Erica and Kate for their comments
on this thesis.
My acknowledgements would not be complete without mentioning the unyielding
support I received from my family and friends. Thank you to my friends at the Thirsty Ear Pub
and in the Microbiology Program. Not only are you brilliant, you’re incredibly fun. My best
friends, Sam, Jackie, Alison, Liz, Sevanne and Danielle, live in NY but have been with me every
step of the way. The laughs and support they have given me have been essential to completing
my Ph.D. To Reid, thank you for inspiring me to work my hardest, for always listening, and for
being so easy to love. To my Aunt Mary Lou who never hesitated to help me when I asked for it,
thank you. And to my parents and sisters, Mom, Dad, Kristen and Tara, your constant
encouragement means so much to me. It is from you that I get my work ethic, my strength, and
my weird sense of humor. I love you all so much and thank you from the bottom of my heart.
5
Table of Contents
Abstract
3
Acknowledgments
5
List of Figures
7
List of Tables
9
Chapter 1
Introduction
10
Chapter 2
Mucin biopolymers prevent bacterial aggregation by retaining
33
cells in the free-swimming state
Chapter 3
Mucins suppress virulence traits of Candida albicans
58
Appendix A
Mucins suppress Pseudomonas aeruginosa virulence toward
83
Candida albicans
Appendix B
Selected antimicrobial essential oils eradicate Pseudomonas spp. and
91
Staphylococcus aureus biofilms
Chapter 4
Conclusions and future directions
6
106
List of Figures
Chapter 1
1-1 Schematic depicting the structure of mucins
13
1-2 Mucins protect the epithelium from colitis and bacterial contact
17
1-3 Mechanisms used by pathogenic microbes to overcome the mucus layer
19
1-4 Models for studying the interactions between microbes and mucus
20
1-5 SEM micrographs of natively purified and industrially purified mucins
22
1-6 Model of mucus protection against pathogenic microbes
24
Chapter 2
2-1 Mucins reduce bacterial biofilm formation
37
2-2 P. aeruginosa swimming velocity is unperturbed by mucins
38
2-3 Nonmotile P. aeruginosa Form Flocs in Mucin Environments
39
2-4 The loss of flagellar motility supports floc formation
40
2-5 Selected viscous polymer solutions support the formation of P. aeruginosa flocs
41
2-6 P. aeruginosa cystic fibrosis clinical isolates form flocs in mucins
42
2-7 P. aeruginosa floc formation in mucins is not dependent on alginate
43
2-8 Flocs grown in mucins are antibiotic resistant
45
Chapter 3
3-1 Mucins induce a unique morphological state characterized by suppressed virulence traits 61
3-2 The mucin-induced morphological state is distinct from the opaque state
64
3-3 qPCR of known C. albicans virulence genes comparing gene expression in RPMI with
65
and without mucins
3-4 The effects of osmotic stress and viscosity on hyphal formation
66
3-5 Mucins suppress hyphae formation in YPD + FBS
67
3-6 Mucins suppress hyphal growth from both yeast and hyphal cells
68
3-7 Mucins reduce attachment of C. albicans to polystyrene and mucus-secreting cells
70
3-8 Mucins reduce C. albicans biofilm formation
72
7
Appendix A
A-1 Mucins protect wild type C. albicans from P. aeruginosa pathogenicity
85
A-2 Mucins reduce P. aeruginosa attachment to C. albicans
87
Appendix B
B-1 Cassia oil kills planktonic bacteria and biofilms with comparable efficiency
94
B-2 Disc diffusion assay identifies essential oils with antimicrobial activity
96
B-3 Activities of selected antibiotics and antimicrobial essential oils against P. aeruginosa
98
PAO1 and P. putida KT2440
B-4 Comparison of two methods of biofilm cultivation for antibiotic and essential oil testing 100
B-5 Susceptibility of S. aureus SC-01 to essential oils
8
101
List of Tables
Chapter1
1-1: Human mucin genes and their expression patterns throughout the body
12
Chapter 2
2-1 List of strains and plasmids used in this study
51
Chapter 3
3-1 Analysis of opaque state attributes upon exposure to mucins
63
Appendix B
B-1 Minimum inhibitory concentration of colistin and essential oils as determined by the
93
standard microbroth dilution assay
B-2 MIC and MBEC of Essential Oil Components Against PAO1
9
102
Chapter 1
Introduction
10
Introduction
The mucus found in the human body covers a vast surface area of epithelial cells that are
exposed to the environment. It coats the respiratory, digestive, and urogenital tracts, as well as
the ocular surface. The mucus barrier is the first line of defense that protects our bodies from
environmental threats including toxins, viruses, bacteria and fungi. However, the mucus layer is
home to a significant portion of the microbiota, or the native population of microbes that inhabit
the body. Despite the enormous microbial population inhabiting the body, many people harbor
these organisms without showing any signs of disease. Certain mucosal infections, such as those
related to inflammatory bowel disease, are accompanied by a disruption of the mucus, suggesting
its importance in keeping infections at bay [1–4]. However, the mechanisms behind mucusmediated protection of the body are not well understood.
In this thesis, I explore the capacity of mucins, the main gel-forming molecules of mucus,
to act as regulators of microbial virulence. Using the bacterium Pseudomonas aeruginosa and the
fungus Candida albicans as model organisms, I determine the effects of mucins on the
physiology of these opportunistic pathogens. My work shows that mucins can suppress microbial
virulence trait expression, highlighting the importance of mucins as regulators of virulence.
Additionally, by combining P. aeruginosa and C. albicans inside a mucin environment, I
investigate the role of mucins in mediating interspecies interactions. Mucins stabilize the
coexistence of these two organisms, suggesting that these polymers not only impact virulence
traits of individual species but can also influence the structure of multispecies microbial
communities.
Mucins are an integral mucus component
Mucus is a mixture of a number of components, including water (~95%), lipids, proteins and
antimicrobial peptides. The slimy, jelly-like consistency of mucus is attributed to mucin
polymers. Mucins are glycoproteins consisting of a protein backbone rich in serine and threonine
residues that are heavily glycosylated, resulting in large (100-10,000kD) molecules. Mucins
polymerize via interacting peptide domains and tangle to form a hydrated mucus gel. The human
body produces cell-surface associated and secreted mucins. Secreted mucins generate the
11
Mucin
Mucin type
MUC1
Membrane
MUC2
MUC3A
MUC3B
Secreted
Membrane
Membrane
MUC4
Membrane
MUC5AC
Secreted
MUC5B
Secreted
MUC6
Secreted
MUC7
Secreted
MUC8
Secreted
MUC9
Secreted
MUC11
Membrane
MUC12
MUC13
Membrane
Membrane
MUC15
Membrane
MUC16
Membrane
MUC17
Membrane
MUC19
Secreted
MUC20
Membrane
MUC21
Membrane
Normal expression pattern
Epithelial surfaces of the respiratory, female reproductive, and
gastrointestinal tracts as well as in the middle ear, salivary, and
mammary glands
Intestinal and colonic goblet cells
Small and large intestine, thymus, liver, lymph nodes, and heart
Small and large intestine, thymus, liver, lymph nodes, and heart
Epithelial surfaces of the eye, oral cavity, middle ear, lachrymal
glands, salivary glands, mammary gland, prostate gland, stomach,
colon, lung, trachea, and female reproductive tract.
Tracheobronchial goblet cells and in the gastric epithelial cells
Salivary, tracheobronchial, and esophageal mucous glands as well
as in the pancreatobiliary and endocervical epithelial cells
Gastric and duodenal mucous glands, pancreatobiliary, and
endocervical epithelial cells
Oral cavity epithelial cells, minor salivary gland, and possibly in
the respiratory tract, pancreas, and bladder
Airway and middle ear epithelial cells and male and female
reproductive tracts.
Female reproductive tract
May represent a differential splice variant of MUC12; Expressed
in the colon, stomach, middle ear, and lung epithelium
Stomach and colon
Gastrointestinal and respiratory tracts
Lung, mammary gland, hematopoietic tissues, gonads, and
gastrointestinal tract
Ocular surface, respiratory tract, and female reproductive tract
epithelia
Gastrointestinal tract with the highest expression in the duodenum
and conjunctival epithelium
Mucosal cells of major salivary glands and the epithelial cells
from corneal, conjunctival, lacrimal gland, middle ear, and
trachea
Highly expressed in the kidneys and moderately in the placenta,
colon, lung, prostate, and liver
Lung, large intestine, thymus, and testis
Table 1: Human mucin genes and their expression patterns throughout the body. Adapted from
[5] with permission from Springer Science and Business Media.
viscoelastic properties of the mucus gel and surface-associated mucins are found in the
glycocalyx, a dense layer of glycoproteins and glycolipids directly associated with the top of the
epithelial layer. Different mucin types are produced in different body regions; there are roughly
20 human mucin genes (Table 1).
12
Figure 1 Schematic depicting the structure of mucins. A) Mucins contain a both nonglycosylated
and glycosylated domains. The nonglycosylated domains are involved in polymerization and
mesh formation. The glycosylated domains allow mucins to retain water to form a hydrogel.
New sketch based on [6] with permission from Elsevier. B) Core glycan structures 1-4 and
examples of mucin glycan structures. Taken from [7] with permission from Cold Spring Harbor
Press.
13
Mucins are produced in goblet cells which synthesize, package, and secrete the polymers
into the environment. For a more detailed description, I refer the reader to a number of
comprehensive reviews [6,8,9]. Briefly, the protein portion of mucins is comprised of several
characteristic domains. The central region contains highly repetitive proline serine threonine
(PTS) domains that are the site of O-glycosylation (Fig. 1A). The N-terminus of cell-surface
mucins contain a transmembrane domain that anchors them to the cell surface. In secreted gel
forming mucins, the C- terminus contains cysteine knot domains and the N or both N and C
termini contain von Willebrand D (VWD) domains, both of which contribute to mucin
oligomerization [10,11].
Secreted mucins are O-glycosylated in the golgi apparatus [12]. O-glycosylation begins
with the addition of an N-acetylgalactosamine residue to a threonine or serine hydroxyl group on
the mucin peptide backbone. Then, galactose and or N-acetylglucosamine residues are added to
form one of four main core structures (Fig. 1B). The core structures can then be further elongated
with sugars including galactose, fucose, and sialic acid, to form glycan chains of varying
complexities. Sialic acids and sulfates are common terminal moieties of mucin glycans, resulting
in a strong negative charge. The composition of mucin glycans is dependent on available
glycosyltransferases which vary throughout the body, thus mucins in different body locations
exhibit different glycosylation patterns. Next, the mucins are either transported to the membrane
or packaged into vesicles for secretion. Once released into the lumen, the mucins quickly absorb
large amounts of water and mix with other components found in the environment to form mature
mucus.
Mucus protects the body from microbial infections
The mucus serves many vital purposes for the body. In the stomach, it modulates pH by
controlling proton transport [13]. In the lungs, the mucus captures particles and microbes from
the environment and is swept away through the coordinated beating of cilia [14]. In the vagina, it
varies in thickness throughout the menstrual cycle to regulate the passage of sperm and prevent
the ascension of bacteria into the uterus [15]. In the intestines, mucus acts as a selective barrier to
allow nutrients to reach the epithelium while preventing microbes from accessing the tissue [12].
While mucus serves specialized roles in each of these regions, it has one common function in all
locations: to protect the underlying epithelia from microbial infection.
14
Microbes are in constant contact with the body. Healthy humans teem with microbes that
inhabit the skin and the mucus. The gut is the most highly populated mucosal region, where
bacterial densities can reach 1011 bacterial cells per gram of feces [16]. Many of these microbes
are commensals that are indispensible for human health. For example, the microbiota is partly
responsible for digestion of food; nutrients that are released as a result of microbial digestion
pass through the mucus and are absorbed by the intestinal epithelium. While pathogens are an
obvious danger to the mucosa, even commensal bacteria can cause disease if they colonize host
cells[17]. Yet despite the high microbial load supported by the mucus, the mucosal epithelium
remains uninfected in healthy people.
Mucin interactions with microbes
The mechanisms of mucus protection against infections are not well understood. One hypothesis
is that mucus acts as a physical barrier to microbes [12]. For example, high viscosity mucus in
the cervix and in the intestines has been shown to decrease the motility of certain bacteria [18].
Mucins play a central role in conferring the physical properties of mucus and are therefore
important determinants of the efficacy of the barrier. Changes in the expression or glycosylation
of mucins can have dramatic effects on the properties of mucus [19]. Additionally, the dense
glycans that protrude from mucins often directly interact with microbes, which can have multiple
sugar binding proteins expressed on the cell surface [20].
Mucins can provide a structural framework for antimicrobial molecules and substances
found within the mucus. They have been shown to bind antimicrobial peptides and may therefore
present them to microbes in high local concentrations. For example, the non-gel-forming
secreted mucin MUC7 binds to the salivary antimicrobial peptide histatin 1, as does MUC5B
which also binds histatins 3 and 5 [21]. In addition to binding these molecules, mucins
themselves can act as antimicrobials. Terminal α1,4-linked N-acetylglucosamine found on
gastric mucins have antimicrobial affects against Helicobacter pylori via the inhibition of cell
wall synthesis [22]. Additionally, MUC7 contains an N-terminal domain with sequence
homology to histatin-5, which demonstrates antimicrobial activity against the fungal pathogen
Candida albicans [23].
Mucins can also bind directly to microbes. Many bacteria are known to associate with
mucins via mucus binding proteins and lectins [24]. For example, the probiotic bacterium
15
Lactobacillus rhamnosus possesses pili that are coated with mucus binding domains [25]. It is
hypothesized that these pili immobilize these bacteria inside the mucus layer, separating the
microbiota from host cells while allowing them to persist as a commensals [26]. Another
possibility is that microbial mucus-binding proteins recognize specific mucin glycans, causing
selective microbial colonization in parts of the body that secrete mucins compatible with specific
microbial binding proteins.
Another potential function of microbial binding to mucins is to serve as decoys for
microbial binding sites to epithelial cells. The microbiota is rich with adhesins and glycosidases
that can adhere to and degrade host-associated glycans, including mucins [27,28]. Since cellsurface mucins can be liberated from the epithelial layer, one hypothesis is that they release upon
binding to microbes, thereby preventing association between host and microbial cells [4].
Additionally, the cell-surface mucin MUC1 reports the attachment of the bacterium P.
aeruginosa to the body via signal transduction, indicating that these mucins behave as signaling
molecules in addition to decoys [29].
Consequences of dysfunctional mucins
The importance of mucus in protecting against microbial infections is demonstrated in disease
states and mucus models in which the mucus layer is disrupted. Mouse models lacking certain
mucins display significant inflammation of the mucosal epithelium. For example, studies using
Muc2 deletion mice show a colitis-like phenotype (Fig. 2A), with inflamed epitheia, bloody
diarrhea and death [30]. In Muc2-/- mice, the commensal bacteria directly contact the epithelium
(Fig. 2B), causing inflammation and cancer development [31]. Additionally, mucus barrier
quality, as measured by penetration by fluorescent beads, is poorer in mouse and human colitis
samples (Fig. 2C&D). However, mucus layer thickness in these samples is not correlated with
permeability, indicating that mucus quality and not quantity is an important characteristic in
inflammatory bowel disease [32]. One explanation for the decrease in mucus barrier effects in
these experiments is that the generation of mucins must be accomplished very quickly during
periods of inflammation, resulting in insufficient concentrations of mucins or mucins with
16
Figure 2 Mucins protect the epithelium from colitis and bacterial contact. A) Muc2 knockout
mice show colitis-like symptoms shortly after birth. Taken from [30] with permission from
Elsevier. B) Mouse colon sections of WT and Muc2-/- mice. Epithelium stained blue and bacteria
stained red. Scale bar: 100 µm. Taken from [31] Copyright (2008) National Academy of
Sciences, USA. C) Control and colitis human colonic biopsy samples inoculated with bacteriasized fluorescent beads on top of the mucus[32]. Mayo 0 represents colitis patients in remission
and Mayo 1-3 represents those with active disease. Scale bar: 100 µm. D) Quantification of bead
penetration through biopsy samples in C) [32]. “Close to epithelium” is <120µm. C&D)
Reproduced from [32] with permission from BMJ Publishing Group Ltd.
17
reduced or abnormal glycosylation patterns that are less effective in preventing disease than
those from healthy people [32].
Mucin glycosylation also plays an indispensible role in conferring the protective
properties of mucus, as evidenced by human disease pathology and studies of glyco-deficient
mouse models. One example illustrating the importance of mucin glycans in human disease is
ulcerative colitis (UC), an inflammatory bowel disease which is linked to altered interactions
between the immune system and the microbiota [33]. Patients with UC have distinct MUC2
glycosylation patterns compared to mucins from healthy individuals [34]. Specifically, UC
patients have a higher abundance of short glycan sequences and fewer long, complex glycans.
Interestingly, non-diseased control groups and UC remission groups show similar glycosylation
patterns, indicating that modified glycans in UC patients reflect their disease state. Abberant
mucin glycosylations in mouse models demonstrate increased disease, and is reviewed
extensively in [35]. Briefly, the effects of the loss of four types of core glycans found on the
intestinal mucin MUC2, namely core-1, 2, 3 & 4 (Fig. 1B), were analyzed in different studies
using glycosyltransferase deficient mice. Mouse models lacking core O-glycans demonstrate
spontaneous development of colitis [36], increased bacterial penetration of the mucus [32] and
increased intestinal permeability and susceptibility to colitis-causing agents [37, 38].
Enhanced susceptibility to disease in the aforementioned glyco-deficient mice may be a
consequence of increased microbial degradation of mucins. Studies of bacterial protease
activities against mucins demonstrate that certain mucin glycans, such as ppGalNAc-T3mediated O-glycosylation, prevent proteolytic digestion of mucins by bacterial proteases [39,40].
Additionally, mucin glycan sulfation and sialylation are hypothesized to decrease or inhibit the
activity of bacterial glycosidases [41, 42]. Therefore, the role of glycans in regulating the barrier
effect of mucus is at least partially due to protection of mucin structure by resisting bacterial
degradation.
Microbial strategies for overcoming the mucus barrier
Certain pathogens have evolved strategies to subvert the mucus barrier to cause infection. These
strategies include physical penetration via pH modulation, enzymatic degradation and avoiding
18
Figure 3 Mechanisms utilized by pathogenic microbes to overcome the mucus layer.
Helicobacter pylori can swim through the normally viscous stomach mucus by increasing its
local pH which reduces mucus viscosity and allows the bacteria to swim through. Other
pathogens, such as Vibrio cholera and Pseudomonas aeruginosa, secrete enzymes that degrade
mucins. The mucus can be avoided entirely by exploiting M Cells, which have little to no mucus
covering. Adapted by permission from Macmillan Publishers Ltd: Nature Reviews Microbiology
[12], copyright (2011)
the mucus by invading M cells, which lack a mucus coating (Fig. 3, [12]). The canonical
example of mucus penetration by a pathogen is H. pylori, which swims through the thick,
viscous mucus of the stomach to infect the epithelium below. This bacterium increases its local
pH, causing the surrounding mucus microenvironment to become pervious through reduction in
viscosity [43]. In addition to raising the local pH, bacteria can degrade mucins to make it more
easily penetrable, as is seen with Vibrio cholera, Yersinia enterocolitica, and Pseudomonas
aeruginosa [39–42]. To avoid the mucus altogether, pathogens can take advantage of Microfold
(M) cells, immune cells that sample the mucosal environment. M cells are found in Peyers
19
Figure 4 Models for studying the interactions between microbes and mucus.
patches in the epithelium, which are regions with little or no mucus that allow for the M cells to
obtain immunogenic materials from the intestinal lumen [48]. These materials are then delivered
to lymphoid tissues, which generate the appropriate immune response. Certain pathogens, such
as Salmonella and Shigella, exploit the lack of mucus on M cells to enter the tissue and cause
infections [49,50].
Models for studying mucus-microbe interactions
A number of model systems for studying mucus are available and vary in their complexity (Fig.
4). The most intricate include mice with mucus irregularities [5] such as mucin deletions [30, 51,
52], mucin overexpression [53], and defective mucin glycosylation patterns [36–38]. These
models are particularly useful in the context of microbe-mucus interaction studies since tissues
can be excised and analyzed to determine the effects of abnormal mucus on the microbial flora.
However, animal models are difficult to maintain and are often insufficient proxies in the context
of microbial disease since many microbes vary in their host specificity. To elude the difficulty in
interpreting animal model data, in vitro and ex vivo techniques are available, such as mucussecreting tissues and cell lines. Sections of mucosal tissues can be excised from humans and
animals and infected with microbes [54–57]. Additionally, cell lines derived from the intestines
and lungs that are capable of secreting mucus are available [58, 59]. These cell lines allow for a
20
controlled environment in which secreted mucus can be harvested for further study or inoculated
with microbes directly. One drawback of these models is that cell culture medium may not allow
for the synthesis of physiologically relevant mucus. Additionally, these cells often have to be
coaxed into mass-producing mucus, which may have reduced barrier effects or altered
characteristics. An alternative model that is particularly useful in human studies is that of whole
mucus. Examples whole human mucus experiments are particularly prevalent in the context of
cystic fibrosis, where patients expel copious sputum that can be collected, and saliva which is
very easily harvested from healthy and diseased people alike [60–63]. Unfortunately, it is
comparatively more difficult to collect mucus from most other body locations, such as the
intestines and the cervix, which require invasive techniques to obtain. Another drawback of
using whole mucus is that there are many components that comprise mucus, making it difficult to
ascertain the specific mechanisms of microbial interactions. Therefore, an even more simplified
model can be used to study mucus-microbe interactions: purified mucins.
To grasp the basics of microbial interactions with mucus, purified native mucins are a
simplified, yet highly informative in vitro system. The glycan structures that coat mucins are
responsible for the physical properties of mucus and provide many potential binding sites for
microbes. Due to the close interactions between microbes and mucins, there is massive potential
to discover strategies employed by these polymers to protect the body against infections. The
purification of mucins enables the in vitro study of mucin-related phenomena without the
confounding factors of whole mucus.
The structure of mucins is crucial for carrying out physiologically relevant experiments,
and high-quality purification of mucins that retains post-translational modifications as well as
supramolecular structure is paramount. In fact, many industrially purified mucins do not form
gels as native mucins do, most likely due to the loss of structural integrity [63, 64] and are
therefore not sufficient comparisons to native mucins. Additionally, the structures of gels formed
by natively purified mucins and industrially purified mucins vary dramatically (Fig. 5), further
highlighting the differences between the two types of molecules.
In this thesis, I use purified native porcine gastric, porcine intestinal, and human salivary
mucins to study microbial interactions with mucin. When human mucins were not available,
such as in in the case of gastric and intestinal mucins, porcine mucins were chosen as a proxy to
21
Figure 5 SEM micrographs of natively purified and industrially purified mucins. Natively
purified mucins show the characteristic fine, heterogeneous mesh structure of mucus, whereas
the industrially purified gel does not.
human mucins due to their high sequence homology [66]. The mucins purified using this method
are structurally and functionally superior to industrially purified mucins because they retain the
important physical and chemical properties that give native mucus its unique characteristics.
Thus, we have the ability to study microbial interactions with mucins in a controlled, highly
reproducible, and physiologically relevant manner. Although purified native mucins closely
reflect the physical properties of mucus, there are a number of limitations to using these mucins
in vitro. Firstly, mucus is made up of numerous components besides mucins, including other
proteins, salts, lipids, antimicrobial peptides and commensal bacteria. While mucins are largely
responsible for the gel forming properties of mucus, the other components are likely to play a
role as well, and may affect microbial behavior. Second, the mucus in the body is constantly
being replenished, which is not reflected in many of the experiments described in this thesis.
Despite the caveats presented here, the use of purified native mucins is highly informative due to
their relative simplicity when compared to other mucus models, their major role in defining the
physical properties of mucus, and their high potential for interacting with microbes. Simplified
22
experiments combining microbes and mucins are an important first step toward enhancing the
understanding of experiments using more complex conditions.
Hypothesis: Mucins protect the body and influence the composition of the microbiota by
suppressing microbial virulence
The examples of microbial interactions with mucus that I have laid out in this introduction are
physical in nature: microbes existing in, binding to, penetrating and degrading mucus. While
these interactions are important in understanding the relationship between the microbiota and the
mucus barrier, these mechanisms do not explain how the mucus prevents these microbes from
causing infection. This thesis aims to explore the influence of mucins on microbial physiology to
understand the mechanisms behind the mucin-mediated protection from microbial disease.
Mucins are emerging as important regulators of microbial behavior that can influence the
expression of virulence traits of microbes. For example, the human cell-surface mucin MUC1
can inhibit surface adhesion of the gastric bacterium Heliobacter pylori [67]. Moreover, the
intestinal pathogen Campylobacter jejuni, influences virulence gene expression in the presence
of human MUC2 [68]. Other examples include modulation of HIV-1 [69] and influenza [70]
infectivity by mucins. However, in-depth analyses of the effects of mucins on virulence
processes and microbial interactions are lacking.
This thesis aims to increase understanding of the influences of mucins on microbes by
performing in depth analyses of bacterial and fungal physiology in the presence of natively
purified mucins. Evidence from the literature demonstrates extensive physical interactions
between microbes and mucins. Therefore, I propose that mucins have direct impacts on the
expression of microbial virulence traits, which functions in tandem with the barrier properties of
mucus to reduce the propensity of pathogens to cause disease (Fig. 6).
Since the human body contains an enormous pool of potential candidate microbes to
study, I began by selecting two well-known model organisms, the bacterium Pseudomonas
aeruginosa and the fungus Candida albicans, to observe their expression of virulence traits in
the presence of mucins. These two organisms were chosen because both P. aeruginosa and C.
albicans can asymptomatically reside in healthy people [71–76] but are formidable pathogens in
the context of disease. Specifically, P. aeruginosa forms robust biofilms that are implicated in
23
Figure 6 Model of mucus protection against pathogenic microbes. (1) The canonical view of
mucus is that it functions as a physical barrier to infection. (2) I propose a second protective
mechanism by which the mucus influences the pathogenic state of microbes: coaxing them
toward commensalism, as opposed to pathogenicity, via the regulation of virulence gene and trait
expression.
wound, eye, and ear infections and perhaps is most well known for infecting the lung mucus of
cystic fibrosis patients. In Appendix B, I show that plant essential oils are more successful in
killing P. aeruginosa biofilms than classic antibiotics. However, it is difficult to deliver essential
oils into the body and there are potential biocompatibility issues, making mucins a more
desirable candidate to study in the context of mucosal surfaces. C. albicans also forms biofilms,
as well as superficial mucosal infections, such as oral thrush and vaginal yeast infections, and
systemic disease. Another reason to study these microbes is that they are typically found near
mucosal surfaces, making them prime candidates for studying in the context of mucins. Finally,
these two organisms have well documented interactions with each other [77–79] and are
therefore logical to study both separately and together inside mucins. Understanding the
influence of mucins on the virulence traits and interactions of these microbes may provide
insight into how they remain in the body without causing disease and may uncover strategies to
prevent them from emerging as pathogens. In the following chapters, I will introduce these
pathogens in more depth and then describe the effects of mucins on them when grown
individually and in coculture.
24
In Chapter 2, I will detail my contributions to a project studying the influences of mucins
on P. aeruginosa biofilm formation [80]. Together with my colleagues, I discovered that mucins
inhibit surface biofilm formation of this pathogen by maintaining cells in a motile, dispersed
state. However, P. aeruginosa can overcome mucin-mediated biofilm inhibition by forming
suspended biofilm-like flocs that resemble those found in the lungs of cystic fibrosis patients.
These flocs are formed not by the wild type, but by nonmotile flagellar mutants, a trait that is
common to cystic fibrosis clinical isolates. Indeed, clinical isolates form flocs in our
experiments, even those that are motile, indicating that other factors besides motility play a role
in floc formation. In Chapter 3, I will detail work aimed at elucidating the influence of native
mucins on C. albicans virulence gene and trait expression [81]. Specifically, I monitored hyphal
formation, surface attachment and biofilm formation in the presence of mucins and found that
mucins suppress them all. Additionally, mucins suppress the expression of a number of
virulence-associated genes, indicating that mucins directly influence microbial physiology. In
Appendix A, the two organisms are combined together to determine the effects of mucins on
microbial interactions. Typically, P. aeruginosa is virulent toward C. albicans, in part by
forming biofilms on fungal hyphae. Since mucins suppress both P. aeruginosa biofilm formation
and C. albicans hyphal formation, I hypothesized that mucins reduce C. albicans killing by P.
aeruginosa. Indeed, mucins extend the viability of C. albicans in coculture, suggesting that
mucin-mediated suppression of virulence trait expression influences microbial community
dynamics.
Overall, the results presented in this thesis suggest that mucins play an important role in
protecting the body from opportunistic pathogens by suppressing the expression of virulence
traits, including aggregation, surface adhesion and biofilm formation. This effect also impacts
microbial community dynamics by suppressing pathogenesis between microbes. I suggest that
mucins act as more than a physical barrier to infection by modulating microbial virulence. In
addition to increasing the understanding of the role of mucins as protectors of the body, this
work demonstrates the potential value of mucins as a strategy for preventing and overcoming
microbial biofouling and infections.
25
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32
Chapter 2
Mucin biopolymers prevent bacterial aggregation by
retaining cells in the free-swimming state
Parts of the work in this chapter were published in:
Caldara M, Friedlander RS, Kavanaugh NL, Aizenberg J, Foster KR, Ribbeck K. 2012.
Mucin Biopolymers Prevent Bacterial Aggregation by Retaining Cells in the FreeSwimming State. Curr. Biol.
Reproduced with permission from Elsevier
33 Abstract
Many species of bacteria form surface-attached communities known as biofilms. Surrounded
in secreted polymers, these aggregates are difficult to prevent and eradicate, posing problems
for medicine and industry [1, 2]. Humans play host to hundreds of trillions of microbes that
live adjacent to our epithelia and are typically able to prevent harmful colonization. Mucus,
the hydrogel overlying all wet epithelia in the body, can prevent bacterial contact with the
underlying tissue. The digestive tract, for example, is lined by a firmly adherent mucus layer
that is typically devoid of bacteria, followed by a second, loosely adherent layer that contains
numerous bacteria [3]. Here, we investigate the role of mucus as a principle arena for hostmicrobe interactions. Using defined in vitro assays, we found that mucin biopolymers, the
main functional constituents of mucus, promote the motility of planktonic bacteria, and
prevent their adhesion to underlying surfaces. The deletion of motility genes, however, allows
Pseudomonas aeruginosa to overcome the dispersive effects of mucus and form suspended
antibiotic-resistant flocs, which mirror the immotile natural isolates found in the cystic
fibrosis lung mucus [4,5]. Mucus may offer new strategies to target bacterial virulence, such
as the design of anti-biofilm coatings for implants.
Introduction
The formation of dense aggregates of cells is key to many bacterial phenotypes, including
those involved in virulence and antibiotic resistance. These communities provide robustness
and protection from environmental threats such as phagocytosis and antimicrobial substances
[6, 7]. The intensive study of surface-attached aggregates, or biofilms, has revealed the roles
of key genes involved in motility, attachment and extracellular polymer secretion [8]. In
addition to the well-studied surface-bound biofilms, many species can also form dense
aggregates in suspension, termed microcolonies or flocs, under certain conditions. The
respiratory tract of cystic fibrosis (CF) patients is a focus for research because the thick static
mucus found in the CF lung leads to chronic colonization by Pseudomonas aeruginosa, which
is a leading cause of morbidity and mortality in CF patients [9, 10]. Successful colonization
by P. aeruginosa results in the formation of flocs (>100µm in diameter) embedded in the
thick mucus layer [11]. In this environment, the bacteria undergo extensive adaptation and
produce virulence factors that altogether assist bacterial infection and inhibit the host immune
defense [12–16].
34 Despite the importance of mucus colonization for bacterial phenotypes and virulence
[11, 17], little is understood regarding the mechanisms that enable efficient growth in mucus,
their native environment. Several observations argue that we cannot generalize from typical in
vitro biofilm studies to the mucosa. For example, clinical P. aeruginosa isolates often
overexpress alginate, an extracellular matrix polysaccharide that is not required for surface
colonization [18]. Additionally, P. aeruginosa frequently switches to the non-motile state
during adaptation in mucus, while motility appears to promote surface attachment during
biofilm formation in vitro [19].
Key to the structural and rheological properties of mucus are mucins—large, densely
glycosylated polymers. We hypothesize that when arranged as a 3D network, pure mucin
polymers can affect P. aeruginosa colonization in the mucosa. Here, we analyze the effects of
mucins on colonization by P. aeruginosa with a simplified mucus model where purified
native mucins are presented in solution. Until now, our understanding of the effects of mucins
on P. aeruginosa has been limited by two key methodological points. First, a direct test of the
effects of mucins on P. aeruginosa aggregation has focused on surface attachment, which is
not required for colony formation. Second, mucins are typically purified using procedures that
degrade and denature the mucin polymers. As we demonstrate below, this results in critical
differences from the native mucus of the lung, where mucin polymers entangle to form a
flexible and highly-organized 3D network.
With our system we show that mucins can effectively suppress bacterial colonization,
both on an immersed solid surface, and in solution. We furthermore show that bacteria can
overcome mucin inhibition and form large flocs upon the loss of flagellar motility. Last, we
show that non-motile, mucus-suspended flocs have superior antibiotic resistance properties
than their motile wild-type counterparts. We conclude that the utilization of a 3D model that
includes native mucins is critical to advance our understanding of mucosal colonization.
Results
Mucins reduce biofilm formation of P. aeruginosa
To begin to dissect mucin-bacterial interactions, we developed an in vitro assay that uses defined
concentrations of native mucins. As a source of mucins we purified native porcine gastric mucus
to obtain an extract composed predominantly of MUC5AC, which is one of the major gel-
35 forming components in the lungs and stomach [20]. The use of natively purified mucins is
decisive for the utility of this assay, as commercially available mucins are processed and have
lost the ability to form viscoelastic hydrogels, as are generated by the native polymers [21, 22].
The second critical feature for this assay is the presentation of mucins in solution, as they exist in
the secreted lung mucus, instead of depositing them onto a surface. This detail is important as the
surface deposition of mucins is likely to adsorb functional groups, thereby partially dehydrating
and altering the biochemical activity of the polymer.
First, we tested the effect of mucins on the ability of bacteria to form biofilms on an
immersed surface. A plastic microcentrifuge tube was inoculated with culture medium that
contained physiological concentrations of mucins [23]. Using the motile, opportunistic pathogen
Pseudomonas aeruginosa, we quantified firm attachment by placing exponential-phase cells into
the mucin solution, allowing biofilm formation to proceed, and quantifying the biofilm and
planktonic populations using the metabolic stain MTT. At 6 h, a time at which biofilms have
begun to form, approximately 90% of P. aeruginosa cells remained planktonic in the presence of
mucins, compared with 50-60% in tryptone broth (TB) alone or TB plus PEG or dextran (Fig.
1A). The total amount of MTT signal between the different conditions illustrates no major
differences in growth, with a small, yet significant, increase in the signal from the mucinexposed population (Fig. 1B). Thus, the results in Fig. 1A are not a result of killing effects by the
mucins. This suggests that mucins suppress biofilm formation, and instead promote a planktonic
lifestyle.
Mucin gels maintain or augment bacterial swimming
It is tempting to speculate that bacteria failed to access the underlying surface because they were
trapped within the mucin network. If this is true, we should expect to see a measurable decrease
of motility within the mucin hydrogel. To test if motion was hindered in the presence of mucins,
we tracked the movements of P. aeruginosa cells that carried a deletion in the flagellar hook
gene (flgE), and were thus deficient in self-propulsion. These cells demonstrated a significant
decrease in diffusivity (p<0.001) in mucin environments, from 2.4 ± 0.2 × 10-9 cm2/s to 1.0 ± 0.1
× 10-9 cm2/s (n ≥ 96 cells), reflecting a higher apparent viscosity of mucin-containing gels, and
suggesting that geometric hindrance was present. However, the wild-type cells remained highly
motile in the presence of the mucins. The distribution of velocities of swimming cells in mucins
36 Figure 1 Mucins reduce bacterial biofilm
formation. PAO1 wild-type bacteria were
grown in polypropylene tubes containing
TB with or without 1% (w/v) PEG,
dextran, or mucin. After 6 h, the amount
of planktonic versus biofilm cells was
quantified using MTT staining. A)
Percentage of the biofilm and planktonic
populations relative to the whole.
Asterisks represent p-value <0.001. Error
bars represent standard deviation of three
replicates. B) Total MTT signal generated
from the combined adherent and
planktonic populations. Asterisk
represents p-value <0.05. Error bars
represent standard deviation of three
replicates.
is similar to that in liquid medium, despite the differences in apparent viscosity (Fig. 2). This
effect was apparent when we compared cells in Pseudomonas minimal medium (PMM) as well
as in tryptone broth (TB) with or without mucins.
Immotile P. aeurginosa cells can form suspended flocs in mucin gels
If mucins can prevent surface colonization by maintaining cellular motility, we speculated that
the loss of cellular motility may be advantageous for colonizing mucus environments. This line
of inquiry may have direct physiological relevance, as isolates of P. aeruginosa from cystic
fibrosis (CF) mucus are often non-motile [5]. Since immotile P. aeruginosa are known to have
reduced surface attachment and biofilm formation, we looked beyond surface adhesion in the
presence of mucin and observed the bacteria in the volume of the mucin gel after 20 h of
incubation. The wild-type cells remained largely as individual cells or small, suspended colonies
of up to 20 µm2 (this corresponds roughly to clusters of 10-20 cells) distributed throughout the
37 Figure 2 P. aeruginosa swimming
velocity is unperturbed by mucins.
Boxplots depicting swimming velocities
of P. aeruginosa in various conditions.
Cells were grown in the media indicated
at
50%-strength.
Velocities
were
obtained from particle tracking analyses
of 20-s swimming videos obtained at 20
frames per second. Data collected by R.
Friedlander.
volume of the mucin medium (Fig. 3A). However, when observing the nonmotile flagellar
mutant PAO1 ∆flgE, we noticed a striking difference compared to the behavior of wild-type
cells. The flagella mutant formed large aggregated flocs of up to 250 µm2 (Fig. 3A). These
differences are not likely due to variations in cellular populations in the mucin medium, as PAO1
displayed similar growth rates in the presence and absence of mucins (Fig. 3B). A similar
behavior was found for two additional flagella mutants, ∆flgK, which lack a hook filament
junction protein, and ∆fliD, which lack an adhesive protein at the tip of the flagellar filament but
not for ∆pilB which lack pilus-mediated adhesion and twitching motility (Fig. 3A). The ability of
cells to form suspended flocs was inversely correlated with their ability to form surface biofilms
in mucin-free environments (Fig. 3C&D). For example, wild-type and ∆pilB cells formed
substantial surface biofilms, but failed to form large suspended flocs in the presence of mucins.
Conversely, the various flagellar mutants formed large flocs, but had reduced surface biofilms in
the absence of mucins. Complementing the flgE deletion in PAO1 ∆flgE restored swimming
motility and diminished the capacity of the bacteria to form flocs in mucin, indicating that it is
indeed the lack of flagella that caused the formation of flocs (Fig. 3E&F).
We hypothesized that loss of flagellar motility (rather than other properties of flagella,
such as adhesion) was the dominant contributor to the observed aggregation. To test this, we
measured mucin-dependent flocculation by a PA14 strain that carries a fully assembled
flagellum, but is paralyzed due to deletions in all four stators in the motor complex
(ΔmotABΔmotCD). This mutant formed substantially larger flocs (up to 60 µm2) than the wild
38 Figure 3 Nonmotile P. aeruginosa Form Flocs in Mucin Environments. A) Floc formation of
PAO1 wild type, flagellar mutants (ΔflgE, ΔflgK, ΔfliD), a pili mutant (ΔpilB), and double
flagella and pili mutant (ΔflgEΔpilB) in PMM with 1% mucins after 20 hr of incubation. Scale
bar is 20 µm. B) Duplication times of the mutant panel in mucins. C) Box plots quantifying floc
size of wild type, flagella, and pili, mutants indicated in µm2 after 20 hr of growth in 1% mucin.
D) Surface-attached biofilm formation of the panel of flagellar and pili mutants. Data are
presented as percent biofilm formation relative to wild type. Error bars represent standard
devation of three replicates. E) Complementation of ΔflgE restores motility and F) suppresses
floc formation. Data in (A-D) collected by M. Caldara.
39 Figure 4 The loss of flagellar motility supports floc formation. P. aeruginosa PA14 without a
flagellum (ΔflgK) and with an immotile flagellum (ΔmotABCD) were inoculated into a 1% mucin
solution in PMM and incubated for 20h before imaging (B-D) and quantification (A). Data
collected by M. Caldara.
type (Fig. 4), but the structures were smaller than those formed by the ΔflgK strain. Both a loss
of motility and loss of the flagella itself, therefore, appear to contribute to mucus colonization.
Notably, floc formation did not occur in PEG, dextran or industrially purified mucins but
did occur in locust bean gum and methylcellulose, two plant-based carbohydrate polymers (Fig.
5). Therefore, not all viscous polymer solutions support the formation of flocs but it is possible
to mimic the effects found in mucins with certain viscous polymer solutions, suggesting that the
physical properties of mucins play a key role in floc formation.
P. aeruginosa cystic fibrosis clinical isolates form flocs in mucins
The loss of flagellar motility allows P. aeruginosa to form flocs in our in vitro mucin gels that
resemble those found in the lungs of cystic fibrosis patients and is also a common characteristic
of cystic fibrosis clinical isolates. Another common feature of clinical isolates is the
overexpression of extracellular polymeric substances, such as the carbohydrates alginate, psl and
40 Figure 5 Selected viscous polymer solutions support the formation of P. aeruginosa flocs. GFP
fluorescent P. aeruginosa PAO1 ΔflgE was inoculated into 0.5% viscous polymer solutions as
indicated on each panel and imaged.
pel. Alginate overexpression, or mucoidy, is particularly common amongst these isolates. We
hypothesized that the mucin gels used in our experiments are a comparable environment to the
cystic fibrosis lung and that clinical isolates form flocs. To test this hypothesis, we inoculated
mucins with four clinical isolates: two that overexpress alginate (FRD1 & 224) and two that
overexpress Psl and Pel (19660 & CF127). The cells were visualized fluorescently by
introducing PBBR1, a plasmid that constitutively expresses GFP using the pLac promoter. All of
the clinical isolates except for 19660 produced large flocs whereas wild-type PAO1 remained
largely individual or formed only small clusters (Fig. 6A). This suggests that elevated secretion
of extracellular polymers strongly promotes the formation of flocs in mucin-environments. The
phenotypes of the suspended colonies resemble those in the cystic fibrosis lung, suggesting that
our mucin-system may present a useful laboratory model for dissecting mechanisms of in vivo
colonization in complex lung mucus.
To validate that the difference in mucus-borne phenotypes is directly due to increased
levels of extracellular matrix, and not a consequence of reduced motility we conducted a
standard motility plate assay [19]. All of the strains except for ΔflgE and FRD1 were motile (Fig.
6B). CF224 and CF127 displayed motility in the plate assay, yet were able to form large flocs in
the presence of mucins. This result indicates immotility is not a requirement for floc formation,
but we cannot rule out the possibility that the contact with mucins induces loss of motility.
41 Figure 6 P. aeruginosa cystic fibrosis clinical isolates form flocs in mucins. A) Fluorescently
labeled cystic fibrosis isolates were grown in 0.5% mucins in PMM for 20h and imaged to
observe floc formation. All strains except 19660 formed flocs. B) Motility assay plates indicate
that all but one isolates are motile.
P. aeruginosa floc formation is influenced by the alginate
Flagellar loss allows bacteria to effectively colonize mucus. Additionally, motile clinical isolates
that overexpress biofilm extracellular matrix polymers display floc formation inside mucin gels.
However, in addition to extracellular matrix overexpression, these isolates may have other
genetic mutations that result in floc formation. Therefore, we tested a panel of isogenic alginate
mutants with varying motility states inside mucins to determine the effects of extracellular
matrix on floc formation. Alginate was chosen because it plays only a minor role in biofilm
formation [24] but is overexpressed in colonies adapted to growth in CF lung mucus [28, 29].
The mutant panel includes an alginate and flagellum deletion mutant strain (ΔflgEΔalgD), an
alginate overexpressing strain (PDO300, a PAO1 strain with a mucoid mucA mutation [27]) and
an overexpressing, nonmotile strain (PDO300 ΔflgE). The singular ΔalgD mutant was not
included in this panel because it does not form flocs, presumably due to wild type levels of
motility. We found that the deletion of alginate in the ΔflgEΔalgD strain significantly reduced
floc size in both 0.5% and 1% mucins when compared to ΔflgE alone (Fig. 7), suggesting that
alginate or its pathway plays a role in floc formation. Interestingly, the motile PDO300 formed
small flocs in 0.5% but not 1% mucins. When the ΔflgE mutation was introduced into the
42 Figure 7 P. aeruginosa floc formation in mucins is not dependent on alginate. A) Fluorescence
micrographs of an alginate mutant panel, both deletions and overexpressors, grown in 0.5% and
1% mucins. B) Floc size measurement for each mutant in 0.5% and 1% mucins. Averages are for
three replicates; Each replicate used five pictures. Asterisks represent p < 0.05. C) CFU counts
demonstrating similar growth patterns of the mutants, therefore any differences depicted in
(A&B) are not due to growth defects. Error bars represent standard deviation of three replicates.
43 alginate overexpressing strain PDO300, floc formation was restored, although the flocs in 0.5%
mucins were significantly smaller than those formed by ΔflgE alone. The restoration of floc
formation in the alginate overexpressing mutant upon flagellar deletion suggests that the loss of
motility is more important for floc formation than the overexpression of alginate. Perhaps at
lower mucin concentrations, P. aeruginosa can overcome the dispersive effects of mucins by
expressing alginate, but at higher mucin concentrations the mucins overcome bacterial
colonization strategies. The decreased floc size in PDO300 ΔflgE is interesting because many
cystic fibrosis isolates overexpress alginate, yet it appears to hinder the formation of colonies as
large as those formed by ΔflgE. Perhaps colony size is less important for bacterial survival in the
body than increased expression of alginate. Alginate overexpression likely provides enhanced
resistance to antibiotics, a phenotype commonly observed in CF pathology [28] and is therefore
highly selected for in the CF lung.
P. aeruginosa flocs that emerge in mucin gels are antibiotic resistant
Last, we asked whether floc formation can provide bacteria with a selective advantage. By
analogy with biofilms, we hypothesized that the immotile cellular aggregates that emerge in
the presence of mucins also have a higher resistance toward antibiotics. We grew wild-type
and non-motile ∆flgE cells in mucin media for 20 h, and then subjected both strains to 20
µg/mL of two clinically relevant antibiotics that differ in their mode of action (Fig. 8). This
experiment revealed two points: first, both wild-type and ΔflgE bacteria were systematically
more resistant to colistin in the presence of mucins as compared to liquid culture without
mucins. This suggests that the mucins themselves have the capacity to reduce the efficacy of
colistin, regardless of whether cells are planktonic (wild type) or form flocs (ΔflgE). Second,
it appeared that the floc-forming ΔflgE cells were more resistant to both antibiotics in the
mucin medium than the motile wild-type cells. To test for this possibility we determined the
percent survival of the bacteria in either condition, by normalizing to the cell numbers in the
untreated samples in liquid and mucin. Inside the mucin medium, the non-motile flagella
mutants were on average 14 times more resistant to colistin (Fig. 8B) and approximately 6
times more resistant to ofloxacin (Fig. 8C) than wild-type cells, both of which are statistically
significant differences. We conclude that the aggregates that emerge upon loss of motility
indeed have an increased resistance compared to motile wild-type cells, possibly due to the 44 Figure 8 Flocs grown in mucins are antibiotic resistant. A) PAO1 Wild type and ΔflgE colony
counts in medium with and without mucins after exposure to the antibiotics colistin and
ofloxacin. B & C) Data from A) depicted as % survival relative to untreated cells. Asterisks
represent different p-value thresholds: (**) if p < 0.01 and (***) if p < 0.001. D) Colony
counts of a panel of PAO1 alginate mutants grown in mucins and exposed to colistin. E) The
data in D) depicted as percent survival relative to untreated cells. Stars represent significant
difference (p <0.05) from the wild type. Error bars in all graphs represent SD of three
replicates.
presence of an altered composition or quantity of extracellular matrix components, or due to a
protective effect of increased cell density [29].
To determine the role of alginate on the resistance of the flocs, we challenged a panel
of isogenic alginate mutants grown in mucins with 20 µg/mL colistin. This experiment was
carried out in 1% mucins, a condition in which only nonmotile strains form flocs (8 D & E).
We found that only the wild type was susceptible to colistin. Both nonmotile, floc forming
strains tested, ΔflgEΔalgD and PDO300ΔflgE were resistant to colistin. The resistance of
ΔflgEΔalgD illustrates that alginate is not not an important component in determining
45 resistance to colistin in the flocs. Interestingly, the motile PDO300 alginate overexpressing
strain was resistant despite the absence of floc formation, suggesting that alginate is sufficient
but not necessary for conferring protection from colistin.
Discussion
Here we have found that animals provide a candidate solution to inhibit biofilm formation,
namely mucin polymers. Critically, our results demonstrate that mucins can limit bacterial
biofilm formation without killing or trapping bacteria, which will help to limit selective
pressure for resistance. Indeed, our only evidence for a resistance phenotype comes in the
form of non-motile cells, which are likely to be strongly limited in other modes of virulence
[5,30]. Our observations of motility and reduced adhesion in mucin media are similar to
findings for Campylobacter jejuni in mouse intestinal crypts. In a previous study, extracted
epithelial scrapings from C. jejuni-colonized gnotobiotic mice demonstrated a lack of
adhesion and unhindered motility within the crypts [31]. Similar to this, a recent study
showed that when supplemented in agar plates, mucins appear to increase motility of P.
aeruginosa [32]. At first sight these and our findings contrast with reports on surfaceimmobilized mucins, which arrest [33,34] and can cause large aggregate formation of P.
aeruginosa cells [35]. However, these findings can be reconciled if one considers that the
effects of mucins on motility may depend on their native three dimensional structure and
hence biophysical properties such as viscoelasticity and lubricity, which are preserved in
native mucus and presumably inside agar gels, but not when adsorbed to a two-dimensional
surface [32]. The gel-forming mucin MUC2 has an ordered repeating ring structure [36], and
we speculate that also other gel-forming mucins, such as the MUC5AC used in our
experiments, display three dimensional features that affect their interactions with bacteria.
Indeed, Berg and Turner have observed that certain structured viscous solutions allow
increased velocities of motile bacteria by providing a rigid framework for generating
propulsive forces [37]. We anticipate that studying mucins in their native three-dimensional
form will reveal valuable novel information about bacterial behavior that cannot be captured
by collapsed mucin monolayers. Indeed, we found that cystic fibrosis clinical isolates form
flocs inside mucin gels that resemble those found in the lungs of cystic fibrosis patients and
that floc-forming bacteria are resistant to antibiotics. Therefore, we suggest that using purified
46 mucins allows for the development of informative in vitro assays for studying the microbial
behavior in mucus.
Experimental Procedures
Mucin purification
The source for purification of native MUC5AC was pig stomachs, which secrete MUC5AC,
homologous to the human glycoprotein [38]. Porcine gastric mucins were purified as described
previously, with the omission of the CsCl density gradient centrifugation [39]. Mass
spectrometry analysis was used to determine the composition of the mucin preparation as
described previously [40]. Briefly, the analysis was performed at the Harvard Microchemistry
and Proteomics Analysis Facility by microcapillary reverse-phase HPLC nanoelectrospray
tandem mass spectrometry on a Thermo LTQ-Orbitrap mass spectrometer. The spectra were
analyzed using the algorithm Sequest [41]. The analysis showed that MUC5AC was the
predominant mucin present in our purified extract, which also contained MUC2, MUC5B and
MUC6 as well as other proteins including histones, actin and albumin. In addition, its quality
was tested by rheology as described in [21,39], which confirmed that the isolated mucins
displayed viscoelastic properties similar to native mucus.
Strains and growth conditions
All strains, plasmids, and their sources are listed below. Pseudomonas aeruginosa PAO1 was the
wild type in this study. P. aeruginosa from the PA14 background was used for the motility
mutants presented in Fig.4. The following media were used: lysogeny broth (LB), tryptone broth
(TB; 10% w/v tryptone), Pseudomonas minimal medium (PMM; 2.5 mM Na-succinate, 1.2 mM
MgSO4, 35 mM K2HPO4, 22 mM KH2PO4, 0.8 mM (NH4)2SO4, E. coli minimal medium (M63
salts) supplemented with 0.2% (w/v) glucose and 0.5% (w/v) casamino acids (M63+). Unless
specified otherwise, the standard growth medium for P. aeruginosa was 1% mucin (w/v) in
PMM. Mucins were dissolved in the medium by gentle shaking overnight at 4°C.
Constructions of deletion mutants and GFP-labeled strains
Deletions in P. aeruginosa strains were obtained using Splicing Over Extension (SOE)-PCR, as
described previously[42], and confirmed by PCR. In addition, the inability of the strains ΔflgE,
47 ΔflgK, ΔfliD to swim and swarm, and the incapacity of ΔpilB to twitch, was tested as described
previously[19]. To facilitate microscopy, we expressed GFP constitutively in each strain. GFP
expressing strains were created as described previously [43]. Strains from the PA14 background
used in Fig. 4 that express GFP were grown in the presence of carbenicillin (250 µg/ml) as
described in [44]. Deletion of fliC was confirmed by PCR and motility agar assay.
Generation of complementation plasmids
A complementation plasmid containing the flgE gene was created using the plasmid
pMQ80[45]. Briefly, the GFP gene was removed from the plasmid via restriction enzyme
digestion with ecoRI and hindIII. The flgE gene was amplified from P. aeruginosa PAO1
genomic DNA with primers containing ecoRI and hindIII restriction sites upstream and
downstream of flgE respectively. After digestion, the flgE gene was inserted into the plasmid
using T4 DNA ligase. For control plasmids, pMQ80 was treated with Klenow polymerase and
blunt end-ligated without an insert. The plasmids were transformed[46] into E. coli DH5α cells
and plated on LB agar containing 30 µg/mL gentamicin selective media. Resulting colonies were
inoculated into overnight cultures and the plasmids were extracted using the GenElute™ Plasmid
Miniprep Kit (Sigma Aldrich). The plasmids were transformed into PAO1 strains plated on LB
agar plates containing 50 µg/mL gentamicin. Successful transformants were again transformed
with the plasmid pSMC21[44], a constitutive GFP expressing plasmid, to facilitate observation
of floc formation (note: the fluorescent strains used in the rest of the study are gentamicin
resistant and could not be used with pMQ80 which relies on gentamicin selection).
Transformants were selected on LB agar containing 400 µg/mL carbenicillin and 50 µg/mL
gentamicin. Floc formation in mucin was performed as described in the main text, with the
addition of antibiotics (carbenicillin for GFP expression and gentamicin for complementation
plasmids) and 100 mM arabinose to induce pMQ80 complementation vector expression.
Quantification of biofilm formation in mucin gels
Freshly growing cells at an OD600 of 0.01 were inoculated in polypropylene PCR tubes and
incubated at 37°C in TB or in TB containing 0.5% (w/v) mucins. After 6 h the planktonic cells
were removed for quantification, and the adherent cells in the tubes were washed 2 times with
PBS to remove non-adherent cells. Planktonic and adherent cells were stained with 5 mg/ml
48 MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) for 2 h at 37°C, and
subsequently destained with 20% sodium dodecyl sulfate in 50% dimethylformamide (adjusted
to pH = 4.7) overnight at 37°C. The resulting solutions were quantified using a plate reader
(OD595).
Particle tracking
For measurement of cell velocities, bacteria were grown to exponential phase as described
above, stained with Syto9 live cell stain by adding Syto9 1:1000 into the culture, and
incubated for 10 minutes at room temperature. The stained cells were diluted 1:10 into a 50%
strength solution of growth medium (as indicated in figure) or growth medium supplemented
with mucin, dextran or PEG. These solutions were mixed and dispensed into chambers for
visualization. Videos of cells were taken on an inverted fluorescent microscope at 20 frames
per second to obtain trajectories (see SI for additional details). The trajectories obtained were
processed using Matlab to determine velocities and diffusivities. Diffusivities were based
upon mean squared displacement values for a range of lag times. Trajectories were also
examined visually to ensure accuracy.
Floc size measurements
To observe cells growing in mucins, cells from exponential growth phase were added to a 0.5%
or 1% mucin gel so that the final concentration was 50-100 cells /µl-1. The same protocol was
used for observing cells grown in other polymeric solutions, specifically, PEG, dextran,
methylcellulose and locust bean gum. The mix was placed in a 96-well glass bottom plate
(MatTek) and incubated at 37°C for 20 h. Images were taken immediately after incubation using
an Axiovert 200M (Zeiss). Floc sizes were quantified in two different ways, by measuring the
size of each floc and by measuring the area of the flocs in each frame of view. In both cases, the
software ImageJ [47] was used. We first subtracted the background using the rolling ball
algorithm. The image was then thresholded using an iterative procedure based on the isodata
algorithm. We defined a group of cells as a floc when it was composed of at least 2-3 cells.
Analysis with Minitab 16 (Minitab Inc.) showed that the data were not normally distributed. We
therefore plotted the data using box-plots to provide an unbiased overview of the distribution of
49 the floc sizes for each condition. Four pictures from three independent experiments were
analyzed for each strain.
Relative biofilm formation
P. aeruginosa Biofilms were grown statically at 37°C on the air-liquid interface of 96-well
plates as described in [48]. The plates were inoculated with tryptone Broth (TB; 1% Tryptone,
0.5% NaCl w/v) containing cells at an optical density at 600 nm (OD600) of 0.0025. After 24
hrs, the biofilms were stained using 1% crystal violet and destained using 33% acetic acid. The
absorbance of the resulting solutions was read with a plate reader at 595 nm.
Motility Assay
M63 motility plates supplemented with 0.2% glucose, 1 mM MgSO4 and 0.5% casamino acids
were created as described previously[49]. To induce expression from the complementation
plasmid, 100 mM arabinose was added to the plates. Wooden inoculation sticks were dipped into
overnight cultures of the strains being tested and used to stab the center of the motility plates.
The plates were incubated overnight (16 h) at 30°C.
Statistical Analysis
Values are reported in the text as value ± SEM. For statistical comparisons between groups with
approximately normal distributions, the Student’s two-tailed t-test was used. Error bars in
figures are either standard deviation or SEM, as indicated in the legends.
Antibiotic treatment
To determine the antibiotic resistance of flocs grown in mucin-media, cells were grown in PMM
with 1% (w/v) mucin. After 20 h, the number of cells was determined by counting CFU; this
number was used as the reference number prior to treatment. The antibiotics ofloxacin and
colistin were added to the cultures at final concentrations of 20 µg/ml, and the cultures were
grown at 37°C for 3 h. After treatment, the number of survivors was estimated by measuring the
CFU. Each experiment was carried out in triplicate. To determine the resistance of cells grown in
the absence of mucins, an exponential phase culture was adjusted to contain the same number of
cells as had grown in 1% mucin in 20 h, and challenged with antibiotics as described above.
50 Acknowledgements
This work was supported by the Cystic Fibrosis Foundation CFF grant number RIBBEC08I0 and
MIT startup funds to KR. KRF is supported by European Research Council grant 242670. RSF is
supported through the National Science Foundation Graduate Research Fellowship Program. We
thank D.J. Wozniak for the EPS deletion strains, B. Berwin for providing the P. aeruginosa
PA14 strains, W. Kim for the labeled conjugating strain, G.A. O’Toole for the complementation
vector, and the lab of Roberto Kolter for the E. coli strain ZK2686.
Table 1 List of strains and plasmids used in this study.
Strains and
Description
plasmids
Reference or
source
E.coli
F- endA1 recA1 galE15 galK16 nupG rpsL
DH10B
ΔlacX74 Φ80lacZΔM15 araD139
Δ(ara,leu)7697 mcrA Δ(mrr-hsdRMS-
Invitrogen
mcrBC) λKmr, thi-1, thr, leu, tonA, lacY, supE,
SM10 λpir
recA::RP4-2-Tc::Mu, pir+ pUX-BF13
[43]
(Apr- Tn7 helper)
S17-1 λpir
Tpr Smr recA, thi, pro, hsdR-M+RP4: 2Tc:Mu: Km Tn7 λpir
Mark Silby
P. aeruginosa
PAO1
wild type, clinical isolate
[50]
ΔflgE
PAO1-ΔflgE
This study
ΔfliD
PAO1-ΔfliD
This study
ΔflgK
PAO1-ΔflgK
This study
ΔpilB
PAO1-ΔpilB
This study
ΔflgE ΔpilB
PAO1-ΔflgE ΔpilB
This study
ΔalgD
PAO1-ΔalgD
51 [51] DJ
Wozniak
ΔalgD ΔflgE
PAO1-ΔalgD ΔflgE
PDO300
PAO1-mucA22 (Alginate overexpressor)
PDO300 ΔflgE
PAO1-mucA22 ΔflgE
This study
PA14
wild type, clinical isolate + pSMC21
[44]
PA14- ΔflgK
PA14-ΔflgK + pSMC21
[44]
PA14- ΔmotAB ΔmotCD + pSMC21
[44]
Cystic Fibrosis Clinical Isolate; Alginate
M. Franklin
overexpressing
[52]
Cystic Fibrosis Clinical Isolate; Alginate
R. Kolter &
overexpressing
K. Foster
Cystic Fibrosis Clinical Isolate; Psl & Pel
M. Parsek
overexpressing
[53]
Cystic Fibrosis Clinical Isolate; Psl & Pel
M. Parsek
overexpressing
[53]
PA14- ΔmotAB
ΔmotCD
FRD1
CF224
19660
CF127
This study
[27]DJ
Wozniak
Plasmids
pMQ30
pBKminiTn7Gm/Cm-gfp
pEX18Gm + CENURA, Gmr, allelic
replacement vector
Gmr, Cmr, transposon delivery plasmid
[54]
[43]
Apr, Kanr, Carbr, plasmid containing GFP
pSMC21
under the control of Ptac constitutive
[44]
promoter
pBBR1_MCS_GFP
Constituitive GFP expression from
pBBR1(MCS5)-Plac-gfp
pMQ80
Complementation plasmid, GmR
pMQflgE
Complementation plasmid containing flgE
52 [55,56]
[45] GA
O’Toole
This study
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57 Chapter 3
Mucins suppress virulence traits of Candida albicans
Work in this chapter was published in:
Kavanaugh NL, Zhang AQ, Nobile CJ, Johnson AD, Ribbeck K. 2014. Mucins Suppress
Virulence Traits of Candida albicans. mBio 5:e01911–14.
58
Abstract
Candida albicans is the most prevalent fungal pathogen of humans, causing a variety of diseases
ranging from superficial mucosal infections to deep-seated systemic invasions. Mucus, the gel
that coats all wet epithelial surfaces, accommodates C. albicans as part of the normal microbiota
where C. albicans resides asymptomatically in healthy humans. Through a series of in vitro
experiments combined with gene expression analysis, we show that mucin biopolymers, the main
gel-forming constituents of mucus, induce a new oval-shaped morphology in C. albicans in
which a range of genes related to adhesion, filamentation, and biofilm formation are downregulated. We also show that corresponding traits are suppressed, rendering C. albicans
incapable of forming biofilms on a range of different synthetic surfaces and human epithelial
cells. Our data suggests that mucins can manipulate C. albicans physiology and we hypothesize
that they are key environmental signals for retaining C. albicans in the host-compatible,
commensal state.
Introduction
Candida albicans is an important opportunistic fungal pathogen in humans that can cause
superficial infections, such as vaginitis in women or thrush in babies and HIV patients, and
systemic, often fatal, disease in more advanced cases [1]. C. albicans possesses a range of
virulence traits, including adherence, filamentation, and secretion of proteases [2]. At the heart of
many infections is the formation of surface-associated C. albicans communities, also termed
biofilms, which can form on mucosal epithelial surfaces and on implanted medical devices, such
as catheters and heart valves. Biofilms show increased resistance to both the immune system and
to antifungal treatment [3]. Despite the ability of C. albicans to cause disease, the healthy human
body accommodates it as part of the microbiota [4–6]. How the body tolerates the continued
presence of potentially virulent C. albicans is largely unknown.
Mucus is the slimy coating found on all wet epithelia in the human body, including the
eyes, airways, and the gastrointestinal and female genitourinary tracts, and many host-microbe
interactions take place in this context. Its major gel-forming components, the mucin
glycopolymers, are emerging as important regulators of microbial virulence. For example, the
human cell-surface mucin MUC1 can inhibit surface adhesion of the gastric bacterium
Heliobacter pylori [7]. Moreover, the secreted human mucin MUC5AC can prevent
59
Pseudomonas aeruginosa surface attachment and biofilm formation by promoting a dispersive
state of bacteria [8]. Other examples include mucus-mediated clearance of the bacterium
Streptococcus pneumoniae [9] and modulation of HIV-1 [10] and influenza [11] infectivity by
mucins. These observations indicate that mucin biopolymers help prevent bacterial and viral
infections by regulating cellular processes related to virulence. Fungal pathogens are only
distantly related to bacteria and viruses, and little is known about their interactions with mucins.
Here, we investigate the role of mucins as potential regulators of C. albicans virulence.
Using a combination of gene expression analysis, mucus-secreting cell lines and defined in vitro
assays with natively purified mucins, we show that exposure to mucins induces a new ovalshaped morphological state in C. albicans, in which various virulence traits are down-regulated,
including surface adhesion, the morphological transition to the filamentous state, and biofilm
formation. Our results indicate that mucins are key contributors to host defense against C.
albicans, and may offer new strategies to target fungal virulence, such as the design of antifungal
treatments or coatings for implants.
Results
Mucins regulate C. albicans physiology
To determine the effects of mucins on C. albicans, the strain SC5314 was cultured in RPMI
medium with and without pig gastric mucins (MUC5AC); RPMI favors growth in the
filamentous state. Mucins were supplemented in the medium to create a 3D environment, as is
found in the native mucus barrier [8]. In the absence of mucins, the cells formed extensive
hyphae which clumped together into flocs (Fig. 1A). In contrast, mucin-exposed cells
predominantly formed short chains resembling pseudohyphae (Fig. 1A) or unicellular, ellipsoidal
cells that are distinct from round, yeast-form cells. An analysis of the growth rate of C. albicans
in mucins shows that cells continue to grow over time, and even show an enhanced increase in
optical density due to the homogenous suspension of individual cells as opposed to the
filamentous flocs formed in RPMI medium alone (Fig. 1B&C).
Importantly, the effect of MUC5AC on C. albicans physiology is not limited to this type
of mucin but was also observed in two other mucins: MUC2 from pig intestinal mucus and
MUC5B from human saliva (Fig. 1D). This indicates that mucins have a general effect on C.
albicans that likely extends across all mucosal surfaces. Due to the abundant availability of
60
Figure 1: Mucins induce a unique morphological state characterized by suppressed virulence
traits A) Phase contrast images of C. albicans in different morphological states. B) Growth curve
of C. albicans +/- mucins in YPD at 30°C. In this case, the growth rates are roughly the same. C)
Growth curve of C. albicans +/- mucin in RPMI at 37°C. Here, the cells grown in mucins
increase in optical density faster than those grown without mucins due to decreased hyphae
formation and flocking of cells in mucins. D) Phase contrast images of C. albicans after growth
in the presence or absence of the following mucins: pig gastric (MUC5AC), pig intestinal
(MUC2) or human salivary (MUC5B).
61
MUC5AC, the remainder of the experiments in this study was performed using this mucin.
At first sight the mucin-induced morphology resembles opaque cells, which are the
mating competent form of C. albicans [12] that have reduced virulence in systemic infection
models [13]. Since the ability of C. albicans to switch to the opaque state is controlled by genes
at the Mating-Type Like (MTL) locus, such that only MTLa or MTLα strains can undergo the
transition to the opaque form, we first tested if mucins induce our starting MTLa/α heterozygous
strains to become homozygous MTLa or MTLα mating-competent cells. Using PCR with primers
specific to the MTL locus, we found that mucin-exposed cells remained heterozygous at the MTL
locus (n = 100). To further probe their identity, the mucin-exposed cells were assayed for three
features that are indicative of opaque cells: the abilities to form opaque colonies on agar plates
[14], mate in the presence of the opposite mating type [15,16], and form mating protrusions in
the presence of mating pheromone [17]. Our data show that the mucin-exposed cells were
incapable of these three traits: they did not form opaque colonies, were not mating competent,
and did not form mating protrusions in the presence of mating α-pheromone (Table 1).
Moreover, exposure of wor1 Δ/Δ cells (which cannot form opaque cells) to mucins also induced
the oval-shaped morphology (Fig. 2), suggesting that the mucin-dependent morphology can
develop independently of the master regulator of opaque status. As one further control, we
repeated the white-opaque switching, quantitative mating and pheromone response assays using
mating competent MTLa or MTLα cells and found that mucins do not affect these opaque
processes in mating competent strains (Table 1). Taken together, these results indicate that
exposure to mucins suppresses the formation of hyphae while inducing the formation of a novel
phenotype that superficially resembles the opaque cell type, which is distinct in its physiological
responses.
Mucins down-regulate virulence-associated genes in C. albicans
To better characterize the mucin-induced morphological change, we carried out transcriptional
profiling experiments. A wild-type C. albicans strain (SC5314) was grown for 8 hrs in RPMI at
37°C in the presence and absence of 0.5% mucins. We performed quantitative PCR (qPCR; Fig.
3) to measure the expression levels of selected virulence genes including those associated with
adhesion [18], biofilm formation [19], and secreted proteinases [20]. TAF145, a general
transcription factor TFIID subunit [21], did not change expression in the presence of mucins and
62
White to Opaque Switch Assay
Strain
Cell Type
Treatment
Switching Frequency
n
SN425
a/α
RPMI + mucin
0%
450
SN425
a/α
RPMI
0%
520
RBY717
a/a white
RPMI + mucin
6.80%
850
RBY717
a/a white
RPMI
5.90%
1200
Response to Alpha Factor
Strain
SN425
SN425
Cell Type
Treatment
a/α
RPMI + mucin
+ α-factor
RPMI + αfactor
RPMI + mucin
+ α-factor
RPMI + αfactor
RPMI + mucin
+ α-factor
RPMI + αfactor
a/α
RBY731
a/a opaque
RBY731
a/a opaque
RBY717
a/a white
RBY717
a/a white
Cells with
Projections
n
0%
200
0%
220
33.80%
330
41.80%
280
0.50%
200
1.00%
200
Quantitative Mating Assay
Strains
Crossed
SN87 X SN95
SN87 X SN95
RBY1177 X
RBY1180
RBY1177 X
RBY1180
Cell Type
Treatment
a/α X a/α
a/α X a/α
a/a opaque X
α/α opaque
a/a opaque X
α/α opaque
RPMI + mucin
RPMI
RPMI + mucin
Mating
Frequency
0%
0%
2.0 %
RPMI
7.9%
Table 1 Analysis of opaque state attributes upon exposure to mucins. C. albicans that is either
homozygous or heterozygous at the mating locus were assayed for their ability to switch from
white to opaque, respond to alpha factor and mate after exposure to mucins.
63
Figure 2 The mucin-induced morphological state is distinct from the opaque state. Phase
contrast images comparing the WT mucin-induced morphology to wor1Δ/Δ, which is locked in
the white phase.
was used as a reference for calibration. This experiment shows that 7 of the 16 tested genes
(indicated by asterisks, Fig. 3) were down-regulated in the presence of mucins by more than 1.5
fold (p <0.05) as determined by a two-tailed, unpaired t-test comparing the ΔCT values of
samples with and without mucins.
Mucins suppress C. albicans transition to the filamentous state
To understand in more detail the effect of mucins on C. albicans physiology, we monitored the
C. albicans strain HGFP3, which expresses GFP from the hyphal-specific HWP1 promoter,
during growth in RPMI with or without 0.5% natively-purified mucins. To test whether the
64
Figure 3 qPCR of known C. albicans virulence genes comparing gene expression in RPMI with
and without mucins. RNA was extracted from 6 independent biological replicates. Error bars
represent SEM. Asterisks represent different p-value thresholds: (*) if p < 0.05, (**) if p < 0.01,
and (***) if p < 0.001.
effects are simply a response to an increased viscosity or osmotic stress, we also subjected the
cells to 0.5% industrially purified mucins (Sigma-Aldrich), 0.5% methylcellulose, or 1M
sorbitol. Industrially purified mucins are proteolytically processed and as a result have lost the
gel-forming capacity characteristic of native mucins [22, 23]. The polysaccharide
methylcellulose is commonly used to mimic the viscosity of a mucus environment [24, 25].
Sorbitol at high concentrations can induce osmotic stress; testing this condition is informative
because the osmotic stress pathway is linked to hyphae formation [26]. Our data show that in the
“no polymer control”, as well as in the presence of methylcellulose or 1M sorbitol, the majority
of cells formed hyphae, as indicated both by the elongated structures and the presence of GFP
fluorescence (Fig. 4). In contrast, in the presence of native mucins, hyphae formation was nearly
completely suppressed. Industrial mucins also inhibited hyphae formation, although not as
effectively as the native mucins; a proportion of cells remained in the hyphal form. These results
suggest that specific biochemical attributes present in the mucins suppress hyphal formation.
Natively purified mucins are obtained by a relatively mild purification procedure to preserve
their structure, and it is formally possible that contaminants contribute to the regulation of the
65
Figure 4 The effects of osmotic stress and viscosity on hyphal formation A) Fluorescent
micrographs overlaid onto phase contrast images of C. albicans grown in RPMI with or without
0.5% mucin (with and without CsCl purification), 0.5% methylcellulose, 1M sorbitol (an osmotic
stress inducer) or 0.5% PEG. The strain used here, HGFP3 fluoresces only when true hyphae are
formed.
C. albicans phenotype. To exclude this possibility, we also tested mucins purified with CsCl, a
more stringent condition which removes the majority of associated proteins and lipids from the
mucins. We observed the same suppression of hyphal formation as in the presence of natively
purified mucins (Fig. 4), indicating that this effect is due to the mucins, and not to any associated
factors.
The ability of mucins to suppress filamentation was also studied at the level of gene
expression. Using qPCR we analyzed the expression levels of the hyphal-specific genes ALS3,
ECE1, and HWP1 in four media conditions: medium without polymers, and medium with native
mucins, industrial mucins, and methylcellulose. For all three genes, native mucins mediated the
strongest down-regulation (Fig. 6B). Of note is that this mucin-induced down-regulation of
hyphal-specific genes was also observed in an alternative culture medium, YPD + FBS (Fig. 5),
indicating that the effects of mucins are not dependent on a specific medium condition.
We next tested the effects of mucins on pre-formed hyphae. Hyphae, formed in the
absence of mucins, were inoculated into RPMI with and without 0.5% mucins or 0.5%
66
Figure 5 Mucins suppress hyphae formation in YPD + FBS. A) Fluorescent micrographs
overlaid onto phase contrast images of C. albicans grown in YPD + FBS, an alternative hyphaeinducing medium to RPMI, with or without 0.5% mucin. B) qPCR comparing the expression of
three hyphal-specific genes with and without mucins in YPD.
methylcellulose. Our data show that newly formed cells, which bud off from the hyphae, were
predominantly in the yeast-form after exposure to mucins (Fig. 6A, right panels). For
comparison, hyphae in the absence of mucins continued to produce hyphae. Hyphal cells
inoculated into medium containing methylcellulose also continued to produce more hyphal cells.
These results were verified by qPCR, which showed that native mucins have the strongest
capacity to suppress hyphal formation (Fig. 6C). We note that the levels of expression of the
gene HWP1 showed a lower degree of suppression (yet significant; p < 0.001) when hyphae were
added as the starting culture in comparison to when yeast were added (-1.3 fold change in
expression with hyphae inoculum vs -15.9 fold change for yeast). Thus, in addition to inhibiting
the growth of hyphae in yeast-form cells, we can also conclude that the hyphal-suppressive
capacity of mucins is strong enough to downregulate the responsible pathway in existing hyphae.
67
Figure 6 Mucins suppress hyphal growth from both yeast and hyphal cells. Fluorescent
microscopy images overlaid on phase contrast images (A) and quantitative PCR of hyphalspecific genes expression from yeast (B) or hyphae (C) after incubation for 8 hours in RPMI ,
0.5% Methylcellulose, 0.5% native mucins or 0.5% industrially purified mucins. The strain used,
HGFP3, expresses GFP only when cells form true hyphae. RNA was extracted from 6
independent biological replicates. Error bars represent SEM. Asterisks represent different pvalue thresholds: (*) if p < 0.05, (**) if p < 0.01, and (***) if p < 0.001.
68
Mucins suppress surface adhesion of C. albicans
A critical, early step of infection by C. albicans is its attachment to a solid surface. In
Fig. 3 and Fig. 6 we observed that genes involved in cell adhesion (ALS1 and ALS3) were downregulated in the presence of mucins. Moreover, we know that mucins may physically trap certain
particles and cells, thereby preventing their association with an underlying surface. Hence, we
tested whether media containing gel-forming mucins also decreases the surface attachment of
C. albicans. We analyzed the ability of C. albicans to colonize two different surfaces in the
presence of mucins, abiotic polystyrene, which is often used in the context of biofilm formation
assays, and human epithelial mucus-secreting cells. For adhesion to polystyrene, a suspension of
yeast cells was inoculated into polystyrene 96-well plates containing RPMI without or with 0.5%
native mucins, industrially purified mucins and methylcellulose. Fig. 7A&B show that native
mucins significantly reduced cell attachment to polystyrene. This effect was detectable as early
as 30 min and became stronger over the course of the hour (unpaired two-tailed t-test; p <0.05).
Methylcellulose was similarly effective in suppressing cell surface attachment (Fig. 7A&B). For
comparison, industrially purified mucins provided no significant protection from surface
attachment. Native mucins and methylcellulose both increase the viscosity of the medium, hence
this parameter could be responsible for the observed anti-adhesion effect. To test this, we
subjected the yeast to medium containing 0.5% of polyethylene glycol (PEG), which is often
used as an antifouling coating [27]. Our data show that PEG was not able to decrease surface
attachment to the same degree as mucins and methylcelluose (Fig. 7C), suggesting that an
increased viscosity alone is not sufficient to protect a surface from C. albicans attachment.
We next infected human mucus secreting cells with C. albicans to determine the effects
of mucus on attachment to a living surface. C. albicans typically resides in the intestine as a
commensal, but this environment can also be the starting point for systemic dissemination [28].
A simple in vitro model for the study of C. albicans interactions with mucus-coated epithelial
cells of the intestines makes use of HT29-MTX cells derived from human colorectal
adenocarcinoma cells, which secrete native-like mucus when growing in culture [29]. The mucus
layer can be removed from the secreting cells with N-acetylcysteine, allowing for a comparison
of cell adhesion with and without native mucus. To determine the effects of mucus on the
attachment of C. albicans to human mucus-secreting epithelial cells, a suspension of C. albicans
was exposed to cells that were lined with an intact mucus layer, or to cells from which the mucus
69
Figure 7 Mucins reduce attachment of C. albicans to polystyrene and mucus-secreting human
cells. (A) Fluorescent microscopy images of polystyrene 96-well plates after incubation with C.
albicans in different media conditions (RPMI alone, RPMI + 0.5% methylcellulose, 0.5%
industrially purified mucin or 0.5% native mucin). Time points were taken every 15 minutes
after removal of non-adherent cells. (B) Quantification of attachment to the polystyrene plates.
Error bars represent standard deviation of 3 replicates. (C) Quantification of attachment to
polystyrene in the presence of viscous polymer solutions. (D) Fluorescence and phase contrast
microscopy images of C. albicans SC5314 stained with calcofluor white after 2 hours of
incubation with HT29-MTX human mucus-secreting cells. (E) Quantification of C. albicans
attachment to HT29-MTX cells. Error bars represent standard deviation of 3 replicates.
had been removed. After 2 hours the unbound cells were removed by washing the epithelial cells
with phosphate buffered saline. Fig. 7D&E show that significantly more C. albicans cells had
attached to the unprotected epithelial cells compared to cells that had been shielded with a layer
70
of mucus. This suggests that the mucus lining of epithelial cells provides efficient protection
from C. albicans attachment.
Mucins decrease C. albicans biofilm formation
C. albicans readily forms surface-attached biofilms on both abiotic and biotic surfaces. Because
biofilm formation relies on attachment, filamentation and cell-cell interactions [30, 31], we
tested if mucins inhibit biofilm formation by C. albicans. We used a standard biofilm assay in
which a polystyrene surface is immersed in a yeast culture (in RPMI) to allow attachment of the
cells, and after 90 min, washed to remove non-adherent cells. Then the surface is submerged in
fresh, cell-free medium to follow the development of the initially attached cells into a biofilm
over 8, 24, and 48 h. We performed this assay in the presence of native mucins and for
comparison, with methylcellulose and industrial mucins.
To evaluate the effect of the individual polymers on C. albicans surface attachment we
removed the supernatant after indicated time points and analyzed the resulting biofilms (Fig. 8).
Surface attachment was visibly reduced in the presence of native mucins, and even more
effectively prevented in the presence of methylcellulose (Fig. 8A). Confocal xy and xz sections
of the biofilms show that both surface coverage (xy images) and thickness of the biofilm are
reduced (xz scan) in the presence of mucins (Fig. 8C). Moreover, a lower percentage of cells in
the mucin-treated biofilms form hyphae within the thin biofilms, confirming the previous
observation from Fig. 4 and Fig. 6. Finally, quantification of the biofilm biomass shows a
significant reduction in the presence of mucins (Fig. 8D). These data suggests that both mucins
and methylcellulose have the capacity to suppress, or destabilize, the attachment of C. albicans
to an underlying polystyrene surface over a relatively long period of time. In Fig. 1B&C we
showed that the cells continue to grow strongly in the presence of mucins, indicating that the
reduced number of attached cells is not due to toxicity of the polymers.
If cell proliferation is not suppressed by the polymers and yet, a reduction of biomass is
observed on the surface, one would expect that a significant proportion of the population has
been shifted to the supernatant. This could occur if cells are released from the biofilm and
continue to propagate in the planktonic phase. Indeed, Fig. 8E shows that the supernatant from
biofilms in the presence of mucins or methylcellulose contained higher numbers of cells than the
supernatants from biofilms growing without mucins. This experiment also reveals another
71
Figure 8 Mucins reduce C. albicans biofilm formation. Biofilms were grown using strain
HGFP3 that produces GFP upon transcription from a hyphal-specific promoter. A) Macroscopic
view of biofilms grown in the presence and absence of mucins or methylcellulose. B)
Fluorescent microscopy images overlaid on phase contrast images of C. albicans found in
biofilm supernatants. C) Confocal images of biofilms grown in the presence and absence of
mucins. D&E) Quantification of biofilm biomass (D) and C. albicans found in biofilm
supernatants (E).
72
important point: in the presence of mucins, the non-attached cells are largely devoid of filaments,
and therefore presumably in a less invasive form (Fig. 8B). In contrast, the cells in the
supernatant of methylcellulose cultures were all highly filamentous. Together these data suggest
that both mucins and methylcellulose can suppress the formation and maturation of biofilms by
reducing attachment of the cells to an underlying surface, possibly by preventing stable
maintenance of cells within the biofilm. However, mucins have a distinct effect: they render
nearly the entire population devoid of filaments.
Discussion
This work shows that the mucin MUC5AC, which is expressed in the stomach and in the lungs,
can induce the downregulation of several virulence traits in C. albicans, both at the level of gene
expression and phenotype. These include the suppression of both filamentous growth and the
formation of surface attached biofilms. Studies with other types of mucins [32, 33], suggest that
the ability of mucins to manage microbial virulence may be a general mechanism that is present
on all mucosal surfaces as part of the innate mucosal immune system.
How might mucins prevent the transition to the hyphal form? Mucins, but not the other
tested polymers, were capable of blocking this transition, suggesting that specific, mucinassociated glycans might be involved in this process. Consistent with this idea, glucose, maltose,
and galactose in solution can all influence the formation of hyphae in C. albicans [34]. The
identity of the mucin glycan moieties that are recognized by C. albicans, as well as the receptors
and pathways in the yeast that are affected by these sugars, are currently unknown; their
identification may suggest new valuable strategies for preventing or recovering C. albicans
infections of mucosal surfaces.
Preventing biofilm formation on materials exposed to living organisms presents a vexing
engineering challenge; the results obtained for mucus hydrogels may provide some interesting
new strategies. Our experiments show that mucins and methylcellulose are both effective in
suppressing C. albicans surface attachment. Both polymers appear to reduce the ability for initial
surface attachment; moreover, they render newly formed cells less capable of stably integrating
into an emerging biofilm. How these polymers work to suppress surface attachment, and even
whether they function by the same mechanisms, are open questions. Despite their superficial
resemblance in surface protection, mucins and methylcellulose have different effects on the
73
surrounding C. albicans cell population. With methylcellulose, the vast majority of cells remain
in the filamentous form, while a cell population exposed to mucins remains largely devoid of
filaments. There is good experimental evidence that the ability to form filaments is required for
C. albicans virulence [35]. Therefore, we have shown that the native mucins in the body are
capable of providing a dual mechanism for virulence control: supressing the yeast-hyphae
transition and surface attachment.
The ability of mucins to suppress virulence traits is not specific for C. albicans but
appears to apply also toward a range of other microorganisms, including the bacteria
Helicobacter pylori and Pseudomonas aeruginosa [7,8] and also certain viruses such as HIV and
influenza [10,11]. Understanding the mechanism of the mucin-Candida host-pathogen
interaction could direct treatment strategies for regulating the healthy microbiota and also shed
light on the molecular origin of increased susceptibility to microbial disease.
Materials and Methods
C. albicans Strains and Media
Strains were maintained on YPD agar (2% Bacto peptone, 2% glucose, 1% yeast extract, 2%
agar) and grown at 30°C. Single colonies were inoculated into YPD broth and grown with
shaking overnight at 30°C prior to each experiment.
The experiments were performed using RPMI 1640 (Gibco 31800-089) buffered with
165mM MOPS and supplemented with 0.2% NaHCO3 and 2% glucose or YPD + 10% Fetal
Bovine Serum (FBS). 0.5% methylcellulose (Sigma, 15 cP) was prepared from a 5% stock
solution by dilution in RPMI. Type II mucin from porcine stomach (Sigma) was dialyzed in a
Spectra/Por Float-A-Lyzer G2 dialysis tube with a 100 kDa molecular weight cutoff, followed by
lyophilization. Industrial PGM and native PGM were dissolved in RPMI and gently vortexed at
4°C overnight.
The C. albicans yeast strains used in this study are SC5314 and HGFP3. Strain HGFP3
[36] was constructed by inserting the GFP gene next to the promoter of HWP1, a gene encoding
a hyphal cell wall protein, and was provided by E. Mylonakis (Massachusetts General Hospital,
Boston, MA, USA) with permission of P. Sundstrom (Stabb et al., 2003).
74
Mucin Purification
The mucins were natively purified to preserve their properties, as opposed to industriallypurified mucins which do not form gels in solution [23,22]. Porcine gastric mucins (PGM) were
purified from fresh pig stomachs as previously described [37]. Briefly, the mucus layer was
isolated from pig stomachs and solubilized in sodium chloride buffer containing protease
inhibitors to prevent mucin degradation, and sodium azide to prevent bacterial proliferation.
Insoluble components were removed via centrifugation, and the mucins were isolated using gel
filtration chromatography on a sepharose column (CL2B). The mucins were then concentrated
and lyophilized. As a control to ensure that there were no contaminants in the mucin preparation,
CsCl gradient centrifugation prepared mucins (as described in [38,39]) were compared to those
prepared without this step.
Growth curves
1 µL of overnight culture was added to 100 µL of medium (RPMI or YPD) in a 96-well plate.
The wells contained plain medium or medium supplemented with 0.5% native mucin, Sigma
PGM or methyl cellulose (Sigma Aldrich, cat # M7140). The plates were incubated at 30°C with
shaking and the optical density at 650nm was read once every hour. The contents of each well
were pipetted up and down before each reading was taken to ensure homogeneous distribution of
cells. After 8 hours of growth, the cells were stained with 10 µg/mL calcofluor white for 5
minutes, aliquoted onto a microscope slide and imaged using a Zeiss Observer Z1 inverted
fluorescence microscope with a Zeiss EC Plan-Neofluar® 20X objective lens (ex365/em445).
Extraction of RNA and cDNA synthesis
1 mL of RPMI or 0.5% PGM in RPMI in a culture tube was inoculated with 10 µL of an
overnight culture of strain SC5314 and incubated at 37°C and 180 rpm for 8 h. RNA was
extracted using the Epicentre® MasterPure™ Yeast RNA Purification Kit and treated with
Sigma-Aldrich® AMPD1 Amplification Grade DNase I. 500 ng of RNA per sample was used to
generate cDNA using the Invitrogen™ Superscript® III First-Strand Synthesis System. cDNA
samples were stored at -80°C until use.
75
Quantitative PCR
All primers were obtained from Sigma and analyzed for efficiency before use in experiments.
Efficiency was calculated by performing qPCR with serial 1:10 dilutions of genomic DNA. BioRad iQ™ SYBR® Green Supermix was used for qPCR reactions. Experiments were performed
in a Roche LightCycler® 480 II machine with the following run protocol: (1) 95°C for 3 min, (2)
40 cycles of 95°C for 10 sec, 58°C for 30 sec, and 72°C for 30 sec. Crossing threshold (CT)
values were obtained and used for analysis. Fold changes were calculated using the ΔΔCT
method in comparison to the reference gene TAF145.
White to Opaque Switch Assays
White-to-opaque switch assays were performed on synthetic dextrose plates as previously
described [40] to determine the white-to-opaque switch frequency for the wild-type a/a white
strain RBY717 [41] after growth in liquid RPMI medium in the presence and absence of 0.5%
purified mucins at 25ºC for 24h. Data is displayed in Table S1. The switching frequency is the
percentage of colonies that displayed opaque sectors or opaque colonies. The “n” is the total
number of colonies counted.
Response to Alpha Factor
The response of C. albicans wild-type white and opaque strains (RBY717a/a and RNY731a/a,
respectively [41] in liquid RPMI medium in the presence of 0.5% purified mucin + α-factor, and
without mucin + α-factor were assayed as described previously [17] after a 24h exposure period
at 25ºC. α-factor in 10% DMSO was added at a final concentration of 10 µg/ml. Cells were
scored for formation of elongated projections 24 h after α-factor was added to the cells. This
experiment was also performed at 37ºC (data not shown), and mucin had no effect on the
observed number of elongated projections in that condition as well. Data is displayed in Table
S1. The “n” is the total number of cells counted, and the ratio of cells with projections was
calculated as a percentage.
Quantitative Mating Assay
Quantitative mating assays were performed as previously described [42] in the presence and
absence of 0.5% purified mucin at 25ºC for 5 days. The wild-type MTL heterozygote strains
76
SN87 and SN95 and opaque strains RBY1177a/a and RBY1180α/α containing different
selectable markers were crossed for this assay. Data is displayed in Table S1. The ratio of cells
with mating products was calculated as a percentage.
Filamentation Assay
100 µL each of RPMI, 0.5% methylcellulose, 0.5% Industrial PGM, and 0.5% native PGM were
inoculated with the strain HGFP3 as yeast-form cells or hyphae in a 96-well plate. 1 µL of an
overnight culture was used as a source for yeast-form cells. For hyphae, an overnight culture was
diluted 1:100 into YPD + 10% FBS, which stimulates the transition to hyphae, and grown to
OD600 = 0.5. The hyphae were spun down, resuspended to OD600 = 5 and 10 µL was added to the
aforementioned conditions. The cells were incubated at 37°C with 180 rpm shaking for 8 h.
Adherent cells were scraped off of the surface and samples were pipetted vigorously to break up
aggregates. 15 µL of each sample was placed on a microscope slide for visualization. Slides were
imaged with a Zeiss Observer Z1 inverted fluorescence microscope with a Zeiss PlanApochromat 20X objective lens under phase contrast and FITC (ex475/em530).
Polystyrene Attachment Assay
Polystyrene 96-well plates were inoculated with 100 µL of RPMI, 0.5% methylcellulose, 0.5%
Industrial PGM, and 0.5% native PGM containing yeast-form cells from the strain SC5314. The
plates were incubated statically at 37°C. Every 15 minutes, a time point was taken by washing
the wells with 200 µL of PBS twice followed by the addition of 100 µL of PBS. After 1 hour, 1
µL of 1mg/mL calcofluor white solution was added to each well. The samples were imaged as
previously mentioned using a 10X objective (ex365/em445). The experiment was performed in
triplicate with 5 pictures taken of each well. The images were analyzed in ImageJ as follows:
each image was converted to 8-bit and the contrast was enhanced (0.4% saturated pixels). The
image was then thresholded to create a binary image. The image was then analyzed using the
“Analyze Particles” tool to measure the surface area covered by cells. The surface area
measurements of the 15 images for each condition and timepoint were averaged.
77
Attachment to Human mucus-secreting colorectal cells (HT29-MTX)
HT29-MTX Mucus-secreting cells reliably secrete a thick, homogeneous layer of mucus as soon
as 7 days post confluency. The cells were grown in a 24-well plate. 2-weeks post confluency, the
cells were treated with 10mM N-acetylcysteine (NAC), which cleaves disulfide bonds between
mucins [43], for 30 min to remove the adherent mucus layer or with PBS as a control. For
infection, C. albicans strain SC5314 was diluted from an overnight culture into DMEM to
OD600 = 0.5. 500 µL of C. albicans was added on top of the HT29-MTX cells and incubated
statically at 37°C for 2 hours. After 2 hours, the medium was removed from the wells, which
were subsequently washed twice with 500 µL of PBS. The remaining C. albicans cells were
stained with calcofluor white and analyzed using a Zeiss Observer Z1 inverted fluorescence
microscope and a plate reader (ex355/em460). The HT29-MTX cells were derived from HT-29
cells (ATCC HTB-38) as described in [29]. HT-29 cells were derived from an anonymous donor.
Biofilm formation assay and Visualization
Biofilms were grown on either of two surfaces: in a 96-well plate (for macroscopic views and
quantification) or on 8mm-diameter-silicone circles (for confocal imaging). Before the
experiment, the silicone circles were washed with water and autoclaved. The surfaces were
incubated in adult bovine serum overnight with shaking at 37°C. The next day, the surfaces were
washed in PBS and submerged in a C. albicans cell suspension of OD600 = 0.5 in RPMI with or
without 0.5% mucins. The samples were incubated at 37°C with shaking at 180 rpm for 90
minutes to facilitate attachment of yeast cells to the surface. Nonadherent cells were washed
away with PBS and the samples were subsequently submerged in fresh RPMI with or without
0.5% mucins, 0.5% industrial mucins, or 0.5% methylcellulose. The biofilms were allowed to
grow with shaking (180 rpm) at 37°C, for the indicated amount of time. For biomass
quantification, the biofilms were stained with 20 µg/mL calcofluor white in PBS for 10 minutes
and analyzed in a Spectra Max M3 plate reader (ex355/em460). Biofilm supernatant
quantification was performed in the same plate reader (absorbance at 600nm). For confocal
imaging, the biofilms were submerged in 6 mL of 20 µg/mL calcofluor white and stained for 10
minutes. The biofilms were imaged using a photo scanner or a Zeiss LSM 700 Upright Confocal.
Planktonic cells were placed on microscope slides and imaged using a Zeiss widefield
fluorescent microscope.
78
Acknowledgments
We thank Dr. Paula Sundstrum and Dr. Eleftherios Mylonakis for strain HGFP3, Dr. Suzanne
Noble for the wor1Δ/Δ strain SN1064, Dr. Richard Bennett for strains RBY717a/a, RBY731a/a,
RBY1177 and RBY1180, and Dr. Thécla Lesuffleur for the HT29-MTX cell line. We also thank
Dr. Bradley Turner for performing CsCl gradient centrifugation. We are grateful to Dr. Gerry
Fink and Dr. Dawn Thompson for helpful comments and advice.
This work was supported by CEHS Pilot Project Grant # P30-ES002109 (K.R. & N.L.K.), the
MIT/NIGMS Biotechnology Training Program Grant # 5T32GM008334-24 (N.L.K.), Burroughs
Wellcome Fund 2012 Collaborative Research Travel Grant (C.J.N.), National Institutes of Health
grant K99AI100896 (C.J.N.) and National Institutes of Health grant R01 AI083311 (A.D.J.).
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42. Bennett RJ, Miller MG, Chua PR, Maxon ME, Johnson AD. Nuclear fusion occurs during
mating in Candida albicans and is dependent on the KAR3 gene. Mol Microbiol. 2005;55:
1046–1059. doi:10.1111/j.1365-2958.2005.04466.x
43. Sheffner AL. The reduction in vitro in viscosity of mucoprotein solutions by a new
mucolytic agent, N-Acetyl-L-Cysteine. Annals of the New York Academy of Sciences.
1963;106: 298–310. doi:10.1111/j.1749-6632.1963.tb16647.x
82
Appendix A
Mucins suppress Pseudomonas aeruginosa virulence
toward Candida albicans
83
Background
The opportunistic fungal pathogen Candida albicans and the bacterium Pseudomonas
aeruginosa are often co-localized in both healthy and diseased humans. As a result of their
coexistence, these two microbes have evolved a pathogenic relationship in which P. aeruginosa
forms biofilms on C. albicans hyphae and secretes small molecules, such as phospholipases and
redox active phenazines, that result in fungal cell death [1, 2]. Although these microbes typically
exist near the mucosa, the impact of mucus on this relationship is unclear. Previous research
from our lab shows that mucins, the main gel-forming components of mucus, suppress virulence
traits of certain microbes, including C. albicans and P. aeruginosa [3, 4]. Therefore, we
hypothesize that mucins likely impact antagonistic interactions between microbes. Through coculturing C. albicans and P. aeruginosa in the presence and absence of mucins, we explored the
effects of mucins on microbial interactions. We found that C. albicans is protected from P.
aeruginosa virulence in the presence of mucins. The protective effects of mucins were lost
toward a constitutively hyphal C. albicans mutant, suggesting that mucin-mediated hyphal
suppression may account for the protective effects. Additionally, we showed that mucins reduce
physical contact between C. albicans and P. aeruginosa, leading to a decrease in bacterial
biofilm formation on the fungus. We propose that through modulation of microbial virulence,
mucins can influence microbial population dynamics and are likely important factors controlling
microbial ecology in more complex communities, such as the microbiota.
Results
We began by coculturing P. aeruginosa PA14 and C. albicans SC5314 in the presence and
absence of porcine gastric mucins mucins (PGM) to determine the effect of the polymers on C.
albicans survival. PGM were chosen due to their enriched MUC5AC content, which is
homologous to human gastric and lung mucins [5]. As a control, C. albicans was grown as a
monoculture with and without mucins to identify any influences on growth; no differences were
observed (Fig. 1A). Next, we added P. aeruginosa PA14 to the experiment to create C. albicans
- P. aeruginosa cocultures. As expected, cocultures grown in the absence of mucins yielded a
reduction in colony forming units (CFUs) as soon as 24 hours after addition of the bacteria, with
complete eradication by 48 hours (Fig. 1A). Interestingly, the addition of mucins delayed
84
Figure 1 Mucins protect wild-type C. albicans from P. aeruginosa pathogenicity. Viability of C.
albicans SC5314 wild type (A) or constitutive hyphal mutant tup1Δ/Δ (B) with and without pig
gastric mucins in monoculture or coculture with P. aeruginosa. (C) Morphology of wild-type
and tup1Δ/Δ C. albicans grown with and without mucins. (D) Viability of wild-type C. albicans
with and without 0.5% methylcellulose in monoculture or coculture with P. aeruginosa.
85
C. albicans eradication by 24hrs, indicating that mucins play protective role against P.
aeruginosa virulence.
One possible explanation for reduced killing of C. albicans in the presence of mucins is
that mucin-induced suppression of hyphae formation protects the yeast from P. aeruginosa
virulence. Previous research demonstrates that P. aeruginosa selectively attached to and kills C.
albicans hyphae, not yeast [1, 2]. To explore this possibility, we tested a constitutively hyphal
strain, tup1Δ/Δ, in the survival assay. This strain forms hyphae within mucins, in contrast to the
wild type in which hyphae formation is suppressed (Fig. 1C). When cocultured with PA14,
tup1Δ/Δ viability decreased at the same rate regardless of the presence of mucins (Fig. 1B),
suggesting that mucin-induced suppression of hyphae formation in the wild type confers the
protection from P. aeruginosa observed in Fig. 1A. Additionally, these results suggest that
mucin protection is conferred by influences on C. albicans and not P. aeruginosa because the
bacterium causes tup1Δ/Δ cell death in both the presence and absence of mucins.
We next tested 0.5% methylcellulose in the survival assay. Methylcellulose is a
polysaccharide that forms viscous solutions that can be used to mimic mucus. It is highly
effective at preventing C. albicans surface attachment, but does not inhibit hyphal formation [3].
This experiment was designed to determine if viscous polymer solutions in general confer
protection, or if suppression of hyphal formation is necessary. During coculture, there appears to
be a small protective effect of methylcellulose at 24 hrs, but by 48 hrs all cells are dead
regardless of the presence or absence of the polymer. This suggests that mucin-mediated
suppression of hyphal formation is more important in conferring protection than the viscosity of
mucins.
One possibility to explain mucin-mediated protection of the wild type is that P.
aeruginosa selectively attaches to and kills hyphae, which are suppressed in the wild-type,
mucin-exposed culture. Another possibility is that a non-attachment mediated form of virulence
is responsible for the selective killing of hyphae, such as secreted factors. To distinguish between
these possibilities, we imaged wild type or tup1Δ/Δ C. albicans with or without mucins, 24, 48
and 72hrs post coculture using scanning electron microscopy. Interestingly, the wild-type,
mucin-exposed C. albicans had fewer attached P. aeruginosa at all time points compared to cells
grown in the absence of mucins (Fig. 2A). To explore this further, the constitutive hyphal
tup1Δ/Δ mutant, which was killed with equal efficiency with and without mucins in the survival
86
Figure 2 Mucins reduce
P. aeruginosa attachment to
C. albicans. Scanning electron
microscopy images of wild-type (A)
and tup1Δ/Δ (B) C. albicans after
coculture with P. aeruginosa PA14
for the indicated period of time.
87
assay, was analyzed using SEM in the presence and absence of mucins. Interestingly, tup1Δ/Δ
showed a slight reduction in attachment at 24 and 48 hrs in the presence of mucins, but was fully
colonized by P. aeruginosa by 72hrs (Fig. 2B). Since the mutant is constitutively hyphal, we
suggest that the presence of mucins and not morphology is the determining factor in bacterial
attachment.
Based on the data from the survival assay, tup1Δ/Δ is killed at the same rate in the
presence or absence of mucins, yet attachment is less prevalent in the presence of mucins.
Therefore, we suggest that killing is not completely dependent on attachment and is partially
dependent on secreted factors. This theory is supported by previous research that shows cell-free
spent coculture medium has killing effects against C. albicans [2]. One possibility is that hyphae
are more sensitive to P. aeruginosa secreted factors and the constitutive presence of hyphae in
the tup1Δ/Δ mutant allows for increased killing by P. aeruginosa despite the anti-attachment
effects of mucins. This also explains the eventual killing of wild-type C. albicans by 72hrs,
despite reduced bacterial attachment.
Discussion
In summary, we showed that mucins had significant impacts on the interactions between C.
albicans and P. aeruginosa. Mucins protected wild-type C. albicans from P. aeruginosa
virulence in coculture, but did not affect a constitutive hyphal mutant, suggesting that
morphology plays a role in determining the protective effects of mucins. However, we found that
mucins suppress bacterial attachment to both the wild type and tup1Δ/Δ, indicting that secreted
bacterial factors are likely more effective in killing C. albicans than attachment. In this model,
hyphae are sensitive to P. aeruginosa derived secreted factors whereas mucin-exposed wild-type
cells are resistant.
While this system suggests a role for mucins as regulators of microbe-microbe dynamics,
it raises the question of how widespread the influences of mucins are. Presumably, a reduction in
microbe-microbe attachment and virulence could have far reaching consequences on the
microbiota. We hypothesize that mucins play large roles in shaping microbial communities and
may be an important determinant of microbiota composition. Since mucins allow for the
coexistence of C. albicans and P. aeruginosa and thereby increase microbial diversity, they
88
could allow for the stabilization of the microbiota and preventing certain species from
dominating the population.
Experimental Procedures
Strains and Growth Conditions
The strains used in this study are P. aeruginosa PA14 and C. albicans SC5314 and tup1Δ/Δ
(courtesy of the Alexander Johnson, UCSF). C. albicans strains were streaked on YPD agar (2%
Bacto peptone, 2% glucose, 1% yeast extract, 2% agar) from glycerol stocks and grown at 30°C.
Single colonies were inoculated into YPD broth and grown with shaking overnight at 30°C prior
to each experiment. P. aeruginosa was inoculated into LB broth from glycerol stocks and
incubated overnight with shaking at 37°C. Experiments were carried out using RPMI (165mM
MOPS, 2% glucose) for C. albicans growth and spent LB (SLB) for coculture. SLB was
obtained by allowing PA14 to grow to OD600 = 1.6, centrifuging the culture and filtering the
supernatant using a 0.2 µm syringe filter.
Coculture survival
1 µL of an SC5314 overnight culture was inoculated into 100 µL of RPMI in a 96-well plate
(Mattek), with or without 0.5% natively purified pig gastric mucins or methylcellulose (15cp,
Sigma Aldrich) and grown for 4 hours with shaking at 37°C. Concurrently, 2mL of LB was
inoculated with 40 µL PA14 and grown for 4 hours with shaking at 37°C. RPMI was then
removed from C. albicans and replaced with 200 µL SLB. P. aeruginosa was added to C.
albicans to a final OD600 = 0.25. A control without P. aeruginosa was included. At 0h, 24h, 48h
and 72h, the contents of the wells were homogenized and a 20µL aliquot was serially diluted in
PBS. 50 µL of the dilutions were plated on YPD agar + 30 µg/mL gentamicin and 60 µg/mL
tetracycline (to select for C. albicans) and Cetrimide agar (to select for P. aeruginosa) and
incubated overnight at 30°C and 37°C respectively. Colonies were enumerated after incubation.
Scanning Electron Microscopy
After coculture for the specified period of time, the cultures were filtered onto 2µm
polycarbonate membranes (Millipore) and fixed with 2% glutaraldehyde. The samples were
equilibrated in 0.1M sodium cacodylate buffer and stained with 1% OsO4. Next, the samples
89
were dehydrated in a series of ethanol baths, critical point dried and sputter coated before
imaging with a JEOL 5600 Scanning Electron Microscope.
References
1.
Hogan DA, Kolter R. Pseudomonas-Candida Interactions: An Ecological Role for
Virulence Factors. Science. 2002;296: 2229–2232. doi:10.1126/science.1070784
2.
Brand A, Barnes JD, Mackenzie KS, Odds FC, Gow NA. Cell wall glycans and soluble
factors determine the interactions between the hyphae of Candida albicans and
Pseudomonas aeruginosa. Fems Microbiol Lett. 2008;287: 48–55. doi:10.1111/j.15746968.2008.01301.x
3.
Kavanaugh NL, Zhang AQ, Nobile CJ, Johnson AD, Ribbeck K. Mucins Suppress
Virulence Traits of Candida albicans. mBio. 2014;5: e01911–14. doi:10.1128/mBio.0191114
4.
Caldara M, Friedlander RS, Kavanaugh NL, Aizenberg J, Foster KR, Ribbeck K. Mucin
Biopolymers Prevent Bacterial Aggregation by Retaining Cells in the Free-Swimming
State. Curr Biol CB. 2012; doi:10.1016/j.cub.2012.10.028
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Turner BS, Bhaskar KR, Hadzopoulou-Cladaras M, Specian RD, LaMont JT. Isolation and
characterization of cDNA clones encoding pig gastric mucin. Biochem J. 1995;308 ( Pt 1):
89–96.
90
Appendix B
Selected antimicrobial essential oils eradicate
Pseudomonas spp. and Staphylococcus aureus biofilms
Work in this chapter was published in:
Kavanaugh NL, Ribbeck K. 2012. Selected Antimicrobial Essential Oils Eradicate Pseudomonas
Spp. and Staphylococcus Aureus Biofilms. Appl. Environ. Microbiol. 78:4057–4061.
91
Abstract
One major challenge posed by cells within biofilms is their higher resistance to antibiotics than
their free-living counterparts. Here, we show that selected antimicrobial essential oils can
eradicate biofilms with higher efficiency than certain important antibiotics, making them
interesting candidates for the treatment of biofilms.
Main Text
Microbial biofilms pose a challenge in clinical and industrial settings where the need for sterility
is paramount. Bacteria within biofilms are more resistant to antibiotics and disinfectants than
individual cells in suspension [1,2]. Several mechanisms can account for the increased antibiotic
resistance in biofilms, including the physical barrier formed by exopolymeric substances [3], a
proportion of dormant bacteria that are inert toward antibiotics [4], and resistance genes that are
uniquely expressed in biofilms [5–8]. Together, these bacterial measures against antibiotics are
urging the discovery of novel strategies that will effectively kill bacterial biofilms.
Plant essential oils have been used for hundreds of years as natural medicines to combat a
multitude of pathogens, including bacteria, fungi and viruses [9]. Several essential oils confer
antimicrobial activity by damaging the cell wall and membrane, leading to cell lysis, leakage of
cell contents, and inhibition of proton motive force [10]. In addition, there is evidence that they
effectively kill bacteria without promoting the acquisition of resistance [11,12]. Finally, many
essential oils are relatively easy to obtain, have low mammalian toxicity, and degrade quickly in
water and soil, making them relatively environmentally friendly [13].
Here, we probe the ability of selected essential oils to kill biofilms formed by
Pseudomonas aeruginosa (PAO1), Pseudomonas putida (KT2440) and Staphylococcus aureus
SC-01. P. aeruginosa is a gram-negative bacterium found in the soil, water, and in animals, but
is also an opportunistic pathogen in humans. It can infect the pulmonary and urinary tracts,
wounds, and burns, and cause devastating medical complications by forming biofilms on medical
devices, such as catheters. The biofilms formed by P. aeruginosa allow this pathogen to evade
treatment with antibiotics and cause persistent, sometimes deadly, infections. The closely related
P. putida can also form biofilms, but is not a pathogen. In rare cases, P. putida can cause
infections in immunocompromised individuals. Usually, P. putida is found in the environment,
especially in soil, freshwater, and on the roots of plants. The gram-positive S. aureus can exist
92
MIC
MIC
P. aeruginosa PAO1 P. putida KT2440
Colistin 3.0 µg ml-1
Not tested
Cassia
0.2% (v/v)
0.2%
Clove
> 5%
> 5%
Lavender
> 5%
> 5%
Peru Balsam
2.5%
2.5%
Red Thyme
> 5%
2.1 ± 0.4%
Tea Tree
5%
2.5%
Table 1 Minimum inhibitory concentration of colistin and essential oils as determined by the
standard microbroth dilution assay. The data here represent the average minimum inhibitory
concentrations of the antibiotic colistin and various essential oils. Each experiment was
performed in triplicate. The highest concentration of each essential oil tested was 5% (v/v). Any
oil that did not show antimicrobial activity in the range tested is listed as “ > 5%.” Standard error
is reported, unless the results for all three trials were identical.
both as a commensal and as a pathogen. As a pathogen, this bacterium is responsible for a broad
range of maladies, from superficial skin infections to serious systemic infections. Treatment of S.
aureus is complicated by antibiotic resistance, which is especially problematic in multidrug
resistant strains such as methicillin-resistant S. aureus (MRSA).
Essential extracts from the bark of plants in the genus Cinnamomum have antibacterial
activity toward a range of different microbes, including P. aeruginosa [14–16]. Importantly, the
effect of Cinnamomum extract on P. aeruginosa was described against individual bacteria in
solution. We then asked if this potent antimicrobial would also be effective against this
bacterium within a biofilm.
To address this question, P. aeruginosa biofilms were grown on the air-liquid interface of
a microscope slide, which was halfway submerged in Mueller Hinton Broth (MHB) containing
PAO1 at an OD600 = 0.0025. After 24 hours of growth at room temperature, biofilms were
93
Figure 1 Cassia oil kills planktonic bacteria and biofilms with comparable efficiency. Cells were
exposed to colistin or cassia oil for 2 h and then stained with a LIVE/DEAD stain to determine
viabililty. Live cells are labeled in green (SYT09), and dead cells are labeled in red (propidium
iodide). Shown here is one representative of three experiments.
washed with H2O and then challenged with cation-adjusted MHB containing 0.2% or 0.1%(v/v)
cassia oil (Cinnamomum aromaticum, 100% pure from Aura Cacia) or 3µg mL-1 colistin. In a
separate assay, the CLSI microbroth dilution method modified with a 2-hour challenge period
[17], 0.2% (v/v) cassia oil and 3µg/mL colistin were determined to be the lowest concentration
of each chemical required to eradicate P. aeruginosa in solution (Table 1). In the case of cassia,
0.1% (v/v) Tween80 was added to mix the oil with the medium [18]. At this concentration,
Tween80 did not affect the growth or viability of planktonic cells or cells in a biofilm (data not
94
shown). After 2 hours, the treated biofilms were rinsed with H2O, stained with LIVE/DEAD®
BacLight™ (Invitrogen), and imaged by wide field fluorescence microscopy. BacLight™ uses a
combination of two nucleic acid dyes: SYTO9, a membrane-permeable green dye that labels
both viable and dead cells, and propidium iodide, a membrane-impermeable red dye that only
labels membrane-compromised cells and eliminates the green SYTO9 signal. Planktonic cells
(final OD600 = 0.25) were challenged with the same concentration of cassia or colistin used
against the biofilms for 2 hours, and then placed into a glass-bottom 96-well plate for imaging.
Our results show that the minimal inhibitory concentration (MIC) of colistin (3µg mL-1)
needed to eradicate planktonic cells is not effective against cells within a biofilm, since a large
fraction of the cells remains stained in green (Fig. 1, top right panel). In contrast, the MIC of
cassia oil against planktonic cells (0.2%, Table 1) is also sufficient to kill the vast majority of P.
aeruginosa cells within a biofilm (Fig. 1, middle panels), suggesting that these cells are not
protected from cassia oil. A slightly lower concentration of the essential oil (0.1%) neither kills
bacteria in solution, nor in biofilms (bottom panels).
Are other antimicrobial essential oils similarly effective as cassia oil in killing
Pseudomonas biofilms? To address this question, we screened for oils that can kill P. aeruginosa
PAO1 in a disc diffusion assay using MHB agar according to the Clinical Laboratory and
Standards Institute standard protocol [19]. The essential oils were supplied by Aura Cacia and
New Directions Aromatics and described as 100% pure. 20µL of each oil was spotted undiluted
onto filter paper discs created from 3-layers of Whatman filter paper (190µm). Our data revealed
the following oils as effective in killing P. aeruginosa: Cassia, Clove (Syzygium aromaticum),
Peru Balsam (Myroxylon balsamum), Red Thyme (Thymus vulgaris) and Tea Tree (Melaleuca
alternifolia) (Fig. 2). To account for the possibility that the oils penetrate into the agar to
different degrees, resulting in what falsely appears to be a reduced antimicrobial effect, any oil
that produced a visible zone of inhibition was considered for subsequent experiments.
In the next step, we explored whether the oils that were active in the disc diffusion assay
are also effective in killing biofilms. To address this point, we determined two parameters for
individual oils: the minimal inhibitory concentration (MIC) required to kill planktonic cells and
the minimal biofilm eradication concentration (MBEC). Biofilms were grown on a MBEC™
device (Innovotech Inc., Edmonton, Canada), a modified microtiter plate that contains 96
polystyrene pegs attached to the lid [1]. The pegs were immersed in MHB containing 106 cells
95
Figure 2 Disc diffusion assay identifies essential oils with antimicrobial activity. Antibiotics at a
concentration of 20 mg ml−1 (A) and pure essential oils (B) were tested against P. aeruginosa
PAO1. The substances that produced a zone of inhibition were further analyzed; lavender oil
served as a negative control.
mL-1 while shaking at 37°C or 30°C for P. aeruginosa PAO1 and P. putida KT2440,
respectively. After 24 hours, the biofilms that had grown on the pegs were rinsed and subjected
to a 1:1 serial dilution of antibiotics and essential oils in cation-adjusted MHB as indicated in
Fig. 3; the medium used to dilute the oils was supplemented with 0.1% Tween80. The volume of
the challenge medium was 200 µl, and the highest concentration of antibiotics and essential oils
tested was 100µg mL-1 and 5%, respectively. Ampicillin and lavender served as negative
controls for antibiotics and essential oils, respectively. After 2 hours of incubation the pegs were
washed, immersed in 150µL fresh MHB, and sonicated for 10 minutes in a Branson 2510
sonicator (40kHz) to release and dissociate the peg-associated biofilms. The average number of
cells on each peg was determined by breaking the pegs off of the lid and sonicating them
individually in microcentrifuge tubes containing 200µL of PBS. The resulting solution was
96
serially diluted and plated onto MHB agar plates to determine colony-forming units. The CFU
counts revealed that the average numbers of cells per peg were 3 × 107 for PAO1 and 4 × 106 for
KT2440. To obtain the MIC, the same number of planktonic cells was added per well to
challenge with antibiotics or essential oils. After 2 hours, 20µL from each well was added to
fresh MHB. After overnight incubation, the lowest concentration of each chemical that prevented
survival the biofilm and planktonic cells was determined. The experiments were performed in
triplicate and the average MIC or MBEC values were determined.
Fig. 3a and 3b show the MIC and MBEC of each chemical tested. Only one antibiotic,
ofloxacin, was able to eradicate planktonic and biofilm bacteria with almost equal efficiency.
The other antibiotics, colistin and gentamicin, were not effective in killing biofilms, even at
concentrations 10-fold higher than the MIC. In contrast, the essential oils cassia and Peru balsam
were effective against biofilms and planktonic bacteria at nearly equal concentrations. This
observation confirms the result in Fig. 1 for cassia oil, where little difference between the MIC
and MBEC was observed. Interestingly, red thyme oil is effective against biofilms at a
concentration of ~2%, but is unable to kill planktonic cells at any of the concentrations tested.
This suggests that thyme oil is more effective against biofilms than it is against bacteria in
solution. To determine statistical significance, we performed a one-sample t-test to compare the
biofilm population to the mean of the planktonic population. Since the planktonic population for
red thyme was not killed by the highest concentration tested, we used the maximum value (5%)
as the population mean to see if significance could be detected at this level. Indeed, the
difference between the planktonic and biofilm populations for both PAO1 and KT2440 were
significant (p <0.05), indicating that red thyme oil is more effective against biofilms than
planktonic cells. We conclude that the essential oils tested here can act against biofilms more
effectively than the tested antibiotics.
To test for potential strain-specific effects of the essential oils we assessed their effect on a close
relative, P. putida (KT2440) (Fig. 3c). Our data illustrate that P. putida is more sensitive than P.
aeruginosa to clove, red thyme and tea tree oils (Fig. 3). This effect is especially evident for
clove oil, which does not eliminate PAO1 but is potent against both KT2440 biofilms and
planktonic cells (Fig. 3). Using one-sample t-tests to compare the P. putida and P. aeruginosa
data for clove oil (assuming that the mean for the P. aeruginosa samples is 5%), we found the
effect of clove oil on P. putida is significantly different from that on P. aeruginosa (P < 0.05).
97
Figure 3 Activities of selected antibiotics and antimicrobial essential oils against P. aeruginosa
PAO1 (A, B) and P. putida KT2440 (C). The MIC and MBEC of various substances were
determined by challenging bacteria that were planktonic or within biofilms, respectively.
Asterisks represent data that extend beyond the plot range, indicating that no killing was
observed at the tested concentrations. Each experiment was performed in triplicate, and the error
bars represent standard error.
98
Additionally, red thyme is effective against planktonic bacteria at 5%, the highest
concentration tested and tea tree oil is effective against biofilm cells. We are unable to calculate
statistical significance in these cases due to the samples surviving at the highest concentration
tested. The differences between the two Pseudomonas species indicate species-specific activity
of the oils and suggest that specific mechanisms of resistance to the oils may be at work. For
example, since certain essential oils appear to work on the cell wall or cell membrane, it is
possible that the composition of these cellular components is key to determining susceptibility to
essential oils. The species-specific activity of the oils suggests that tailored combinations to
target a range of different microbes may be effective against multi-species biofilms.
One similarity between the P. aeruginosa and P. putida data is that red thyme oil is more
effective against biofilm cells than their planktonic counterparts. The same is true for tea tree oil
against P. putida, which is ineffective at concentrations of 5% or less against planktonic bacteria,
but is effective against biofilm bacteria at a concentration of ~4%. In these cases, being inside a
biofilm turns into a disadvantage to the bacteria, as it renders them more susceptible to the
activity of this particular essential oil. It is possible that the extracellular matrix of the biofilm
adsorbs the active components and increases their local concentration. Another possibility is that
the cell membrane or cell wall in biofilm cells is different than planktonic cells due to differential
gene expression in the two cell types.
It should be noted that the data obtained using the MBEC device is reproducible with a
different assay where biofilms are grown in the wells of a 96-well microtiter plate [20] instead of
on polystyrene pegs (Fig. 4). In this assay, the protocol is the same as that for the MBEC device,
with the difference that the plates are not shaken during incubation, allowing the biofilms to
grow on the sides of the wells at the air-liquid interface. Additionally, the biofilms are formed in
TB (1% tryptone, 0.5% NaCl) as opposed to MHB. In Fig. 4, we compare the susceptibility of
biofilms that were grown with both approaches. The biofilms grown with the two different
methods contain comparable number of cells (2x107 CFU/well vs. 3 x 107 CFU/peg). The MBEC
values obtained for each method were the same for all substances except ofloxacin. It is unclear
why cells grown on the MBEC device are more susceptible to ofloxacin than cells grown in 96well plates, especially considering that the other substances tested do not show significant
variation in efficacy. However, it is possible that the difference in the medium or structure of the
biofilms caused the increased susceptibility of biofilms grown on the MBEC device to ofloxacin.
99
Conc. of antibiotic (μg/mL)
A 100
Conc. of essential oil (% v/v)
* *
*
* *
90
80
70
60
Wells
50
MBEC
Device
40
30
20
10
0
B
* *
Colistin
Gentamicin
Ofloxacin
* *
5
Ampicilin
* *
* *
4
3
2
1
0
Cassia
Clove
Peru
Balsam
Red
Thyme
Tea Tree Lavender
Figure 4 Comparison of two methods of biofilm cultivation for antibiotic and essential oil
testing. P. aeruginosa biofilms were grown either on the sides of wells in a 96-well plate or on
the pegs of an MBEC device, and their sensitivities toward antibiotics and essential oils were
determined.
After testing two closely related gram-negative bacteria, we studied the effect of essential
oils against the gram-positive bacterium S. aureus (Fig. 5). Our goal was to determine if the oils
discriminate between gram-positive and gram-negative bacteria. The strain used in this study,
SC-01, is a biofilm-forming, oxacillin- and methicillin-resistant clinical isolate [21]. Certain
essential oils, such as tea tree, thyme, and peppermint, are effective against planktonic [18,22,23]
and biofilm [24,25] MRSA. However, essential oils from cassia, red thyme, or clove have not
100
Figure 5 Susceptibility of S. aureus SC-01 to essential oils. (A) A disc diffusion assay reveals
that SC-01 is sensitive to various essential oils. (B) The MIC and MBEC of essential oils were
determined by challenging planktonic cells and biofilms, respectively. Asterisks represent data
that extend beyond the plot range, indicating that no killing was observed at the tested
concentrations. Each experiment was performed in triplicate, and the error bars represent errors.
been tested against MRSA biofilms of any strain. Moreover, the strain used in this study (SC-01)
has not been challenged with essential oils in previous work. First we performed a disc diffusion
assay to determine if the same oils that are effective against Pseudomonas work against S.
aureus. Indeed, all of the oils tested, including lavender, showed a zone of inhibition (Fig. 5A).
101
MIC
Cinnamaldehyde 0.1% (v/v)
MBEC
0.2%
Eugenol
>5%
3.3 ± 0.8%
Linalool
> 5%
> 5%
Table 2 MIC and MBEC of Essential Oil Components Against PAO1
Next, we tested the essential oils against biofilms formed on the pegs of the MBEC
device. The protocol is the same as described above, and the average number of cells per peg
was 1.5 x 105. The results show that the biofilms were killed by the same or similar
concentrations of cassia, Peru balsam, and red thyme oils as were effective against P. aeruginosa
(Fig. 5B). Notably, this strain is resistant to methicillin yet is killed effectively by four essential
oils tested in this assay.
After determining that essential oils are effective against biofilms, we tested individual
components of the essential oils for antimicrobial efficacy. We assessed the molecules
cinnamaldehyde, eugenol and linalool (from cassia, clove and lavender oils, respectively) [26,27]
for their effect against P. aeruginosa planktonic and biofilm cells. All three components were
obtained from Sigma Aldrich. The protocol used in this assay is identical to that used for testing
whole essential oils, including the use of Tween80 in the medium to suspend the components,
which have a low solubility in water. Table 2 summarizes the data, which indicate that
cinnamaldehyde is as effective as the complex cassia oil. Additionally, whereas clove oil is not
effective in killing P. aeruginosa biofilms in 5% v/v solutions, its ingredient eugenol is effective
at 3.3%. The finding that single essential oil components are effective at eradicating bacterial
biofilms is promising, as it may allow the dissection of their mechanisms of action, as well as
inspire the molecular design of new antimicrobial components.
In summary, we demonstrate here that the essential oils cassia, Peru balsam and red
thyme are more effective in eradicating Pseudomonas and S. aureus biofilms than selected
important antibiotics, making them interesting candidates for the treatment of biofilms.
Important future goals include identifying further active antimicrobial components within the
oils, as well as the molecular mechanisms by which these components so effectively breach the
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biofilm barrier. In this study, we only sampled a small number of different oils, but a plethora of
other oils is available in nature, bearing an enormous potential for the discovery of alternative
treatments to antibiotics.
Acknowledgements
We thank Marina Caldara for discussion of intermediate results and for valuable comments on
the manuscript. This work was supported by the MIT startup funds for Katharina Ribbeck.
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Chapter 4
Conclusions and future directions
106
In this thesis I propose that mucins defend the body from infections by modulating
bacterial and fungal virulence and supporting microbial diversity. I found that mucins effectively
suppress virulence traits in two opportunistic pathogens: Pseudomonas aeruginosa and Candida
albicans. The mucin-mediated effects on virulence not only impact each microbe individually,
but also the relationship between the two when grown in coculture. Specifically, mucins reduce
P. aeruginosa pathogenicity towards C. albicans, indicating that mucins can affect the microbial
composition of cocultures. Based on this work, I hypothesize that mucins have considerable
potential to influence human health by modulating bacterial physiology and diversity in the host
microbiota.
Prior to this thesis, few studies focused on the effects of mucus on microbial physiology;
most research concentrated on the opposite effect where microbes impact mucus integrity and
expression. While such studies are indispensible for understanding the barrier properties of
mucus during mucosal diseases, the influence of the mucus environment on microbial
physiology is equally important. Characterizing mucus-mediated effects on microbes allows us
to elucidate effective strategies for manipulating microbial physiology, particularly in ways that
reduce virulence.
In Chapter 2, I showed that mucins suppress biofilm formation of wild type P.
aeruginosa. Biofilms are highly problematic due to their mechanical robustness and significant
resistance to antibiotics. Plant essential oils are also effective at targeting bacterial biofilms when
antibiotics fail, as I show in Appendix B. However essential oils can have toxic effects at high
concentrations inside the human body and are difficult to deliver to some parts of the mucosa,
such as the lungs and intestines. Therefore, mucins may serve as effective, biocompatible
materials to reduce biofilm formation. The mechanisms by which mucins suppress biofilm
formation are unknown, but several possibilities exist. Mucins may form surface coatings that
prevent bacterial adhesion. Indeed, previous studies show that mucin coatings spontaneously
form on a number of surfaces and repel binding of mammalian [1] and bacterial cells [2].
Another possibility is that mucins occlude P. aeruginosa surface proteins that are involved in
surface attachment. For example, P. aeruginosa binds to mucins via the flagellum [3–5], which
is also involved in adhesion to surfaces [6]. Mucins may therefore block flagellar surface
attachment domains either by binding to them directly or by steric hindrance.
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Although mucins are successful in inhibiting surface associated biofilms, we show that
P. aeruginosa can overcome the protective effects of mucins when it loses flagellar motility. The
loss of motility is a common characteristic of P. aeruginosa cystic fibrosis isolates, consistent
with this being advantageous for the colonization of mucus [7]. In our experiments, flagellar
deletion mutants form flocs that resemble biofilms in their structure and demonstrate resistance
to antibiotics, much like the colonies formed in the lungs of cystic fibrosis patients. Indeed,
clinical cystic fibrosis isolates also formed flocs in mucin solutions. Interestingly, the loss of
motility is not a requirement for floc formation, since a number of motile clinical isolates also
colonized mucus. Future work includes elucidating the traits of the clinical isolates that allow
them to colonize mucus despite their motility. Regardless of the mechanism, the ability to
recapitulate clinical phenotypes in vitro presents a model system in which P. aeruginosa clinical
isolates can be studied in a physiologically relevant manner. For example, bacteria from cystic
fibrosis patients could be inoculated into mucins and challenged with antibiotics to more
accurately predict successful treatment options.
Although P. aeruginosa is able to overcome the suppressive effects of mucins through
the loss of motility, the dispersive effect of these polymers on wild-type bacteria suggests that
they have potential to reduce microbial virulence. Indeed, mucin-mediated virulence trait
suppression is also observed with the fungus C. albicans. In Chapter 3, I found that mucins
suppress the yeast to hyphal transition, surface attachment, biofilm formation, and the expression
of a number of virulence-associated genes in C. albicans. The modulation of virulenceassociated gene expression by mucins hints at an exciting potential function of these polymers: to
persuade microbes to remain in an avirulent state when they are in close proximity to the
epithelium.
The suppression of a selection of C. albicans virulence traits by mucins suggest that these
polymers are important for protection from disease but a number of open questions remain. First,
do mucins impact non-virulent processes? Presumably, the answer is yes. C. albicans is highly
responsive to environmental factors such as mechanical stress, sugar concentration and oxygen
availability [8–10], therefore high concentrations of mucin glycans and the mucin hydrogel
structure likely impact many aspects of C. albicans physiology. Secondly, how do mucins
modulate virulence gene and trait expression? The work presented in this thesis does not
elucidate the mechanisms by which mucins suppress C. albicans virulence traits, but future work
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to address this point may include studies of global gene expression. Only a small portion of
virulence-related genes were analyzed in this study. Therefore a broader characterization of the
impact of mucins on gene expression may suggest the involvement of previously characterized
pathways, which would enhance our understanding of mucin-mediated effects on C. albicans
physiology. Another method to guide the discovery of mucin-mediated virulence suppression is a
deletion mutant screen. For example, a library of characterized C. albicans deletion mutants
could be inoculated into mucins and observed for the formation of hyphae in the presence of
mucins, which is normally suppressed. Although such a study may provide valuable insights into
the mechanisms of mucin-mediated virulence trait suppression, one caveat regarding these types
of studies is that C. albicans has redundancies in many of its pathways, including filamentation
and adhesion [11–13], making it difficult to find single deletion mutants that behave differently
from the wild-type in the presence of mucins. Screening a panel of transcription factor mutants
allows for the identification of gene networks involved in the response to mucins, and may
alleviate concerns of redundancies if redundant genes have common regulatory networks.
Regardless of these caveats, the use of global gene expression analysis or mutant libraries is a
first step that can be followed by more focused approaches, such as the analysis of deletion
mutants or the study of single proteins.
In Appendix A, P. aeruginosa and C. albicans were combined inside mucin
environments to determine the impacts of the biopolymers on microbial community dynamics. P.
aeruginosa is antagonistic toward C. albicans, typically outcompeting the fungus within 48
hours of coculture. However, the presence of mucins allowed the microbes to coexist for 72
hours. In this system, mucins influence microbial diversity by reducing bacterial virulence
toward its fungal counterpart. The coculture experiments suggest that the suppression of
microbial pathogenicity induced by mucins is beneficial for microbes by allowing them to thrive
in environments that are normally deadly. In this scenario, mucins not only impact microbial
virulence, but also the structure of the microbiota, which is emerging as an important
determinant of human health. Certain microbiota compositions in humans are linked to different
health and disease states, such as obesity [14], inflammatory bowel disease [15] and arthritis
[16]. One can imagine that people with mucin defects, such as aberrant glycosylation patterns or
insufficient mucin production, may experience dysbiosis that leads to disease. This idea is
supported by a study in which a mouse strain deficient in the B4galnt2 glycosyltransferase,
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which glycosylates intestinal epithelial cell surfaces, shows alterations in the intestinal
microbiota composition compared to wild-type mice [17], suggesting that glycans are important
determinants of the microbiota composition. An analysis of mucin glycosylation patterns may
therefore provide insight into patient health, both to diagnose and to predict ailments. This idea
has been put forth in the context of ulcerative colitis, where patients with active disease show
altered mucin glycosylation patterns when compared to healthy individuals [18].
Supplementation with healthy mucins may prove to be therapeutic for those suffering from
dysbiosis related to insufficient or damaged mucins.
The use of mucins as therapeutics has many advantages, one being that they suppress
microbial virulence without sacrificing cell viability. The reduction in P. aeruginosa and C.
albicans biofilm formation without cell death is particularly important, because antibiofilm
methods that are bacteriocidal, e.g. antibiotics or disinfectants, often lead to the development of
resistance over time. In the case of mucins, the selection of resistant population by killing
sensitive cells does not occur, therefore reducing the risk of evolved resistance. Evolution assays
in which microbes are passaged daily into fresh mucins can be employed to test if the inoculum
loses its sensitivity to mucins over time. For example, the C. albicans strain HGFP3, which
expresses GFP only when true hyphae are formed, could be monitored for fluorescence after
each passage to determine if mucins lose their ability to suppress hyphae formation. Mutants
would then be sequenced to identify evolved loci and thereby the mechanism used by the fungus
to gain resistance.
Important to note is that none of the other tested polymers, namely industrial mucins,
methylcellulose, or PEG, display all of the effects of mucins. In the case of C. albicans,
industrial mucins suppressed hyphal formation but did not disrupt surface attachment. The
opposite case was true for methylcellulose, which reduced surface attachment but not hyphal
formation. Mucins can confer multiple methods of virulence trait suppression against C.
albicans, which suggests that they are highly evolved to tackle many facets of microbial
virulence. In the coculture assay, methylcellulose did not extend the viability of C. albicans as
was seen with mucins. Perhaps the presence of both peptides and glycans components on
mucins, both of which can vary based on gene expression and bodily location, allows for finely
tuned influences on microbial behavior that cannot be achieved by the presence of a viscous
solution alone. Because the manipulation of mucin glycan structures is very difficult to achieve
110
in a standardized manner, thorough analysis of mucin structure may elucidate important mucin
characteristics that influence disease. For example, molecular characterization of glycan structure
and the peptide backbone can be performed on mucins from patients that suffer from C. albicans
infections and compared to those from healthy people.
To strengthen the idea that mucins are general regulators of microbial physiology and
community structure, a range of microbes need to be studied in the presence of mucins, including
pathogenic and commensal bacteria and fungi. An example from one of my colleagues shows
that mucins reduce biofilm formation of the Streptococcus mutans, a bacterium responsible for
dental cavity formation [19], suggesting that these polymers have broad effects on different
microbial species. Finally, more complex but perhaps more physiologically relevant studies of
dual species or multi species communities in mucus are needed to determine the effects of
mucins on microbial community structure.
The work presented in this thesis indicates that mucins influence microbial virulence
traits and interactions, which represents a new perspective on the protective functions of mucus.
Mucus is typically considered a physical barrier to infection. Here, I show that mucins have
massive potential to reduce microbial virulence and impact the composition of the microbiota. A
picture of the far-reaching influences of mucins is just starting to emerge, and our work already
suggests that these polymers are widely effective against microbial virulence. This work
represents a first step toward understanding the potential of mucins with the hope that one day
we can harness their strategies to develop effective antimicrobial therapeutics.
111
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