IAI Accepts, published online ahead of print on 25 August 2014 Infect. Immun. doi:10.1128/IAI.01876-14 Copyright © 2014, American Society for Microbiology. All Rights Reserved.
1
Localization of Burkholderia cepacia Complex Bacteria in
2
Cystic Fibrosis Lungs and Interactions with Pseudomonas aeruginosa
3
in Hypoxic Mucus
4 5
Ute Schwab1*, Lubna H. Abdullah1, Olivia S. Perlmutt1, Daniel Albert2, C. William Davis1,
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Roland R. Arnold3, James R. Yankaskas1, Peter Gilligan4, Heiner Neubauer5, Scott H. Randell1**,
7
and Richard C. Boucher1**
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Cystic Fibrosis/Pulmonary Research and Treatment and Research Center1, Marine Sciences2,
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School of Dentistry3, Pathology and Laboratory Medicine, Memorial Hospital4 , University of
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North Carolina, Chapel Hill, USA; Friedrich-Loeffler-Institut, Institute of Bacterial Infections
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and Zoonoses, Naumburger Str. 96a, D-07743 Jena, Germany5.
12 13
Running title: Burkholderia cepacia complex bacteria in the CF lung
14 15
* Corresponding author, present address:
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Clinical Sciences-Immunology and Microbiology
17
College of Veterinary Medicine
18
Cornell University
19
Ithaca, NY 14853
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Phone 607-229-2399
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Email: [email protected]
22
** Senior authors contributed equally to this work.
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This work is dedicated to the memory of our friend and colleague Dr. Anthony Paradiso. 1
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ABSTRACT
25
The localization of Burkholderia cepacia complex (Bcc) bacteria in the cystic fibrosis (CF) lung,
26
alone or during co-infection with Pseudomonas (P.) aeruginosa, is poorly understood. We
27
performed immunohistochemistry for Bcc and P. aeruginosa on 21 co-/or singly-infected CF
28
lungs obtained at transplantation or autopsy. Parallel in vitro experiments examined growth of
29
two Bcc species, Burkholderia cenocepacia and Burkholderia multivorans, in environments
30
similar to those occupied by P. aeruginosa in the CF lung. Bcc was predominantly identified in
31
the CF lung as single cells or small clusters within phagocytes and mucus, but not as “biofilm-
32
like structures”. In contrast, P. aeruginosa was identified in biofilm-like masses, but densities
33
appeared to be reduced during co-infection with Bcc. Based on chemical analyses of CF and
34
non-CF respiratory secretions, a test medium was defined to study Bcc growth and interactions
35
with P. aeruginosa in an environment mimicking the CF lung. When test medium was
36
supplemented under anaerobic conditions with alternative electron acceptors, B. cenocepacia and
37
B. multivorans used fermentation rather than anaerobic respiration to gain energy, consistent
38
with the identification of fermentation products by HPLC. Both Bcc species also expressed
39
mucinases that produced carbon sources from mucins for growth. In the presence of P.
40
aeruginosa in vitro, both Bcc species grew anaerobically, but not aerobically. We propose that
41
Bcc: 1) invades a P. aeruginosa infected CF lung when the airway lumen is anaerobic; 2)
42
inhibits P. aeruginosa biofilm-like growth; and 3) expands the host bacterial niche from mucus
43
to also include macrophages.
44
2
45
INTRODUCTION
46
Individuals with cystic fibrosis (CF) typically exhibit polymicrobial lung infections, with
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P. aeruginosa becoming highly prevalent as the disease progresses (1). A subset of CF patients
48
become infected with Burkholderia cepacia complex (Bcc) bacteria (2). Bcc infected CF patients
49
exhibit significantly reduced long-term survival (3), reflecting both a more rapid decline in lung
50
function and, in some cases, the “Cepacia syndrome”, characterized by unrelenting pneumonia,
51
sepsis, and death.
52
However, there is a paucity of information on the localization of Bcc alone, or in
53
conjunction with P. aeruginosa, in the CF lung. Based on the observation that both pathogens
54
can be isolated from CF sputum samples, it is generally believed that Bcc and P. aeruginosa co-
55
localize in macro-colonies (“biofilm-like structures”) within the lungs of chronically infected CF
56
patients. This notion has been supported by in vitro studies demonstrating mixed biofilms in
57
vitro (4-6) and bacterial use of the same chemical signals to control biofilm formation and
58
expression of virulence factors (5). It has also been shown that growth inhibitory substances
59
produced by P. aeruginosa can protect their biofilms against invasion by Bcc, but when no
60
inhibitory components are produced, the two species are able to co-exist (7).
61
There is evidence, however, that these organisms may not co-exist in biofilm-like
62
structures within the CF lung. For example, P. aeruginosa has been shown to grow in sessile
63
biofilm matrices, both in vivo and in artificial sputum medium mimicking the CF lung habitat (8-
64
14). In contrast, Bcc biofilm-like structures have not yet been identified within the intraluminal
65
CF mucus inhabited by P. aeruginosa. The absence of mixed biofilms in excised lung
66
specimens is perplexing because Bcc isolates have been shown to form biofilms in vitro,
67
regardless of species (15, 16), and we previously reported the formation of biofilm-like
3
68
structures on the surface of well-differentiated (WD) human airway epithelial cultures (17).
69
Therefore, it is unclear whether Bcc biofilms in vivo have been missed, or that the current in
70
vitro systems that generate biofilms do not adequately mimic the CF lung environment.
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Other data suggest that P. aeruginosa and Bcc may occupy different niches within the CF
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lung. Several clinical and laboratory observations suggest that the niche occupied by Bcc in the
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CF lung is within pulmonary phagocytes and respiratory epithelial cells rather than in
74
intraluminal mucus. These observations include clinical unresponsiveness of Bcc to
75
antimicrobial therapy despite an isolate’s susceptibility in vitro, isolation of serum-sensitive
76
isolates in bacteremic infection (18), and the close taxonomic relationship between Bcc and the
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intracellular pathogen B. pseudomallei (19). Intracellular survival of Bcc is reported in multiple
78
pulmonary cell types, including a transformed human bronchial epithelial cell line from a CF
79
patient (20), CFTR-positive and CFTR-negative bronchial epithelial cell lines (21), the
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transformed human bronchial epithelial cell line BEAS-2B (22), polarized lung epithelial cells
81
(23), A549 pulmonary epithelial cells (24, 25), primary type II pneumocytes (26), and pulmonary
82
macrophages (27-30). These data are complemented by a clinical and pathologic study that
83
localized Bcc within macrophages in freshly excised CF lungs (31). However, because the
84
observations of Sajjan et al. (31) were restricted to CF lungs infected by the transmissible clonal
85
ET 12 strain of B. cenocepacia (Edinburgh-Toronto) lineage (genomovar III, cblA+) (32), it is
86
not known whether the immunohistologic findings represent characteristic features of this
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species/strain or late stage pulmonary infection with Bcc in general. Importantly, the
88
localization of P. aeruginosa was not investigated in CF lungs from which sputum evidence for
89
P. aeruginosa and Bcc co-infection existed.
90
Consequently, we designed a study to localize both Bcc and P. aeruginosa in CF lungs,
4
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excised at transplantation or autopsy, infected with one or both organisms. We also designed
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parallel in vitro experiments to ask whether Bcc could survive in environments similar to those
93
occupied by P. aeruginosa in the CF lung. When mucus plugs and plaques exceed a certain size,
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and are adjacent to highly metabolic cells, the intraluminal CF mucus can become markedly
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anaerobic (10). Thus, the anaerobic physiology of P. aeruginosa has been studied extensively
96
(10, 33-37). However, the literature on energy production by Bcc in hypoxic environments is
97
sparse. Consequently, we also performed experiments using airway surface layer (ASL) material
98
harvested from WD airway epithelial cultures as a relevant medium to study how Bcc gains
99
energy under O2-deprived conditions, and the interspecies relationships between Bcc and P.
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aeruginosa during growth under anaerobic and aerobic conditions.
101 102
MATERIALS AND METHODS
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CF patients and specimens. Sputum and lung tissue specimens were obtained from 21 CF
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patients under University of North Carolina (UNC) Institutional Review Board (IRB) approved
105
protocols consistent with all federal and institutional requirements for informed consent and
106
confidentiality. Ten were female and eleven were male (age range 13 to 48 years), 18 lungs
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were obtained at lung transplantation and 3 at autopsy (Table 1). Tissues were dissected to
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isolate bronchial and bronchiolar specimens that were fixed in neutral buffered formalin,
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embedded in paraffin, and sectioned using conventional procedures.
110
Sputum samples were obtained prior to specimen procurement, consistent with routine
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patient care. Samples obtained closest to the surgical/autopsy dates were analyzed for the
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presence of Bcc, P. aeruginosa and other bacteria at the University of North Carolina Hospital
113
Clinical Microbiology Laboratory. When possible, the relative bacterial densities were
5
114
determined by a dilution streak plate method. For this semi-quantitative method, a grade of 0,
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1+, 2+, 3+, or 4+ was assigned depending on the growth seen in each dilution streak on a plate
116
(0=no bacteria and 4+=bacteria present after three dilution streaks).
117
We selected 2-4 tissue blocks from each lung based upon airway size and the presence of
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mucus within the airway lumen. When possible, two blocks with large airways and 2 blocks
119
with small airways and alveoli were selected (Table S1).
120
Rabbit antisera. Hyperimmune antisera were produced by immunizing rabbits with inactivated
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whole-cell Bcc or P. aeruginosa antigens and were stored at -20oC until further use. Specificity
122
of the antisera was determined by Western blot against crude bacterial cell lysates, including 5
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CF P. aeruginosa isolates and 5 Bcc strains (i.e., B. cepacia ATCC 25416, B. cenocepacia, B.
124
vietnamiensis, B. multivorans, and B. dolosa). Bacteria for lysates were grown in TSB, washed in
125
PBS, and adjusted to an optical density at 600 nm of 1 (equal to ~5 × 109 bacteria/ml). Then, 2
126
ml of the whole bacteria were centrifuged for 10 minutes at 5000 X G. The pellet was lysed (125
127
mM TrisHCl pH 7.0, 5.0 mM NaF, 2 mM EDTA, 1% NP-40, 1 mM Na3VO4, 100 μM TPCK,
128
100 μM quercetin, 1 mM PMSF, 1 μg/ml leupeptin, and 1 μg/ml pepstatin, 2% SDS, 20%
129
Glycerol), boiled for 5 minutes at 100°C, sonicated for 15 seconds, and aliquots were stored at -
130
20°C. For Western blots, 10 µl per lane of bacterial lysates were run on NuPage Bis-Tris gels
131
(Invitrogen) and electrophoretically transferred to a polyvinylidene fluoride membrane. The
132
membrane was blocked with 5% powdered milk in Tris-buffered saline with 0.1% Tween-20
133
(TBST) for 1 hour. Blots were incubated with antisera at 1:50,000 and 1:100,000 dilutions in
134
blocking solution overnight at 4°C, washed with TBST, and incubated for 1 hour with
135
peroxidase goat-anti-rabbit secondary antibody diluted 1:100,000 in blocking solution. After
6
136
washing with TBST, signal was detected with Immobilon Western Detection Reagent (Millipore)
137
and exposure to film. A duplicate gel was silver stained to evaluate loading.
138
Immunohistochemistry. Bcc and P. aeruginosa were localized in formalin fixed paraffin
139
sections of CF lung tissue obtained during transplantation or autopsy (Table 1 and 2). In almost
140
all cases, 4 tissue blocks were selected from each patient, 2 containing larger bronchi with
141
cartilage in the airway wall and 2 containing bronchioles, in which cartilage was absent and
142
alveoli were present. Sections were deparaffinized, rehydrated, and subjected to antigen retrieval
143
by boiling in 0.05 M Na citrate pH 6.0. Endogenous peroxidase was inhibited by incubation in
144
3% H2O2 in methanol, sections permeabilized with 0.1% Triton X-100 (in PBS), and washed in
145
PBS, 0.05% Tween 20 (PBST). After blocking in 5% donkey serum, 1% fish gelatin, 1% BSA
146
in PBST, sections were incubated overnight at 4oC with 1:3000 and 1:6000 dilutions of P.
147
aeruginosa or Bcc antisera, respectively. Optimal dilutions for each primary antibody were
148
determined using positive control sections containing easily identifiable Pseudomonas colonies
149
(from a CF patient infected only with P. aeruginosa; Table 1, patient #21) or numerous intra-
150
and extra-cellular B. cenocepacia bacteria (autopsy tissue from a Cepacia syndrome CF patient,
151
Table 1, patient #1). These positive control sections were included in each staining run.
152
Negative controls consisted of double the concentration of normal rabbit serum in place of the
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primary antibody on replicate sections of every tissue block. After washing with diluent (1:3
154
blocking solution in PBST), sections were incubated with biotinylated Fab fragment donkey anti-
155
rabbit secondary antibody (Jackson ImmunoResearch, West Grove, PA), washed in diluent, and
156
incubated with streptavidin horseradish peroxidase (Jackson ImmunoResearch). Slides were
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washed with PBST, rinsed in 0.05 M Tris-HCl buffer, pH 7.6 and incubated with
158
diaminobenzidine tetrahydrochloride (10 mg/50 ml) in the same buffer with 0.006% H2O2. After
7
159
rinsing in water, sections were counterstained with ethyl green, dehydrated in ethanol, and
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coverslipped in Permount. A Nikon microphot SA microscope connected to a 3CC-Chilled
161
Camera (Sony, Marietta, GA) interfaced to a computer was used to capture light microscopic
162
images via Adobe Photoshop.
163
Slides were evaluated by two independent observers and were graded as strong (+++),
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moderate (++), weak (+) staining for P. aeruginosa and/or Bcc in endobronchial mucus and
165
other locations. Each slide was examined for bacterial appearance (single rods, clusters, or
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macrocolonies), location of bacteria (inside inflammatory, bronchial epithelial or alveolar cells)
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and morphology of airway epithelium (intact, partial or total denudation).
168
Supernatants of mucopurulent material (SMM) and airway secretions obtained from CF
169
and non-CF patients. Under UNC IRB approved protocols, mucopurulent respiratory secretions
170
were harvested from CF lungs excised during transplantation (female, n=10; male, n=10; average
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age, 29.2 years), and from donor lungs that were unused for transplantation (non-CF control;
172
female, n=6; male, n=4; average age, 32.6 years). Approximately 1-5 ml of mucopurulent
173
material (CF) or pooled secretions (control) was directly aspirated from intrapulmonary airways
174
with syringes, ultracentrifuged for 1 h at 100,000 rpm (434,902 x g, Beckman TL-100 Tabletop
175
Ultracentrifuge, TLA 100.2 rotor), and the supernatants were stored at -20oC prior to analyses
176
(38, 39).
177
In vitro Production of Airway Surface Layer (ASL) materials. Because ASL is a relevant
178
culture medium for our studies, we developed techniques to harvest large quantities of ASL from
179
WD airway epithelial cell cultures prepared as previously described (40). Cells were removed
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from portions of main stem or lobar bronchi from excess donor tissue obtained at the time of
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lung transplantation or from CF lung tissue, again under UNC IRB approved protocols. Cells
8
182
were grown on 100-mm tissue culture dishes in bronchial epithelial growth medium (BEGM).
183
Passage two cells were seeded on collagen IV coated 30 mm Millicell CM membrane supports
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(Millipore) at a density of ~106 `cells/cm2 and grown in air-liquid interface (ALI) media in the
185
absence of antibiotics. Upon reaching confluence (~5 days), the culture medium was only
186
replaced in the serosal chamber. At ~21 days, when the apical cell surface of cultures was fully
187
differentiated into ciliated and mucus producing cells (as determined by light microscopy), 200
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l of Kreb’s Bicarbonate Ringer (KBR) solution was deposited onto the apical cell surface
189
followed by collection of ASL 24 h later with a pipette tip attached to a 3 cc syringe.
190
Preliminary experiments revealed that KBR was conditioned by the epithelium to resemble ASL
191
in vivo with respect to ionic composition within 6 to 12 h (41). After ASL collection, the apical
192
cell surface of cultures was washed with endotoxin free PBS and, after 24 hours, KBR was added
193
for a second harvest of ASL. The harvested ASL was stored at 4oC.
194
Chemical analysis of SMM and in vitro produced ASL. The ions Na+, K+, Cl-, and Mg2+ were
195
measured using the Vitros950/950AT Chemistry System by standard protocols. The electron
196
acceptors nitrate and sulfate were measured in secretions and in vitro ASL by ion
197
chromatography using a Dionex 2010i Ion Chromatograph. Test samples and standards were
198
diluted 1:50 in sodium bicarbonate/sodium carbonate eluent, filtered (0.45 µm filters) to remove
199
particulates, and 1 ml of sample injected onto the columns. Standards were run to determine
200
NO3- and SO42- peaks, sample areas then compared to standards, a standard curve graphed, and
201
the sample mass calculated according to the slope. Iron concentrations were measured
202
spectrophotometrically using a commercial kit (MPR Iron without deproteinisation, Roche
203
Diagnostics Corp., Indianapolis). Amino acids were measured by HPLC (Millipore, Waters 7R
204
WISP). Osmolality was measured by using a Micro osmometer Model 3300 (Advanced
9
205
Instruments, Inc.). The data obtained from in vivo secretions and ASL measurements were used
206
to develop an artificial nutrient limited medium that was used to study bacterial growth in a
207
defined, pathophysiologically-relevant microbiology system. Thus, the range of growth rates
208
produced are predicted to be much slower than those obtained using nutrient-rich laboratory
209
media (e.g. Luria Bertani medium) and may provide a more accurate indication of Bcc bacterial
210
physiology relevant to the CF lung. P. aeruginosa growth and biofilm formation have already
211
been studied in synthetic media that nutritionally mimic CF sputum (11, 12, 42).
212
Purification of mucins. Commercially available mucins are poorly defined chemically and non-
213
gel forming. MUC5AC and MUC5B are the major gel-forming mucins in both bovine cervical
214
mucus and human respiratory mucus (43-45). To produce large volumes of purified
215
Muc5AC/5B mucins, 50 mL of fresh bovine cervical mucus was solubilized in 6 M guanidinium
216
chloide and mucins were purified by double isopycnic density gradient centrifugation (46).
217
Fractions were collected, weighed for density, and assessed for DNA (A260), protein (A270), and
218
sialic acid. Sialic acid-positive fractions were pooled and dialyzed against deionized water. The
219
second gradient centrifugation yielded a sharp sialic acid peak that was well separated from
220
DNA. A known volume of this fraction was dried and its weight determined on a microbalance
221
(CAHN, C33) to determine the mucin content. Mucin solutions, reconstituted with PBS at
222
concentrations ranging from 0.01 to 1 mg in a volume of 100L, were prepared and used for
223
bacterial growth studies.
224
Bacterial strains. Two Bcc species, B. cenocepacia BC-7 (genomovar III, cblA+ major CF
225
lineage ET12; provided by Dr. Richard Goldstein, Boston University) and B. multivorans J-1
226
(genomovar II; UNC Hospitals) and the P. aeruginosa strains PAO1 (ATCC) and Pae 33 (UNC
227
Hospitals) were used in this study. Pae 33 was from an individual with severe lung disease being
10
228
evaluated for lung transplantation. The patient had two additional mucoid strains of
229
Pseudomonas and this smooth strain was resistant to many antibiotics, suggesting long standing
230
infection. Stocks of the bacteria were kept at –70oC in skim milk. For all studies, bacteria were
231
grown at 37oC on sheep blood agar plates.
232
Inoculation of in vitro produced ASL and “pure” mucin solutions. To characterize the
233
growth of B. cenocepacia BC-7 and B. multivorans J-1 in ASL produced in vitro and in mucin
234
solutions, we incubated s small volume of ASL or mucin solution (30 µl) with a low number,
235
(approximately 200 to 500 cfu/1 μl PBS) of bacteria (grown overnight on sheep blood agar and
236
then suspended/diluted in PBS). The cultures were grown for 3 days and their growth
237
monitored under aerobic and anaerobic (10% H2, 5% CO2, 85% N2, anaerobic chamber from
238
Coy Laboratory Products, Inc., Ann Arbor, MI) conditions at 37oC in round bottom 96 well
239
plates. All growth studies were performed in triplicate. For anaerobic growth experiments, both
240
ASL and bacterial suspensions were equilibrated in the anaerobic chamber before ASL was
241
inoculated with Bcc and/or P. aeruginosa. The number of viable bacteria was determined by
242
serial dilutions plated onto sheep blood agar and cultured under aerobic conditions.
243
To test mixed cultures for growth in ASL under aerobic/anaerobic conditions, equal
244
numbers of P. aeruginosa (approximately 200 cfu/0.5 μl PBS) and Bcc (approximately 200
245
cfu/0.5 μl PBS) as well as a different ratio of Bcc to P. aeruginosa ( 100:1 were co-cultured, and
246
after 72 hours, serial dilutions were prepared and plated in parallel onto trypticase soy agar and
247
Bcc selective agar (provided by the Microbiology/ Immunology Laboratory of UNC Hospitals)
248
for bacterial enumeration.
249
Preparation of bacterially conditioned test medium and in vitro ASL supernatants. In vitro
250
ASL was inoculated with the two Bcc species and P. aeruginosa bacteria as described above in
11
251
quadruplicate, and after 72 h of incubation, the four samples were pooled, filtered (0.2 µm
252
Acrodisk) and ASL filtrates stored at –20oC until use.
253
Measurements of organic acids in infected test medium and in vitro ASL. For these studies,
254
bacteria-conditioned and non-inoculated test medium and ASL were assayed for low-molecular
255
weight organic acids, e.g., acetate, lactate, or formate. These acids are fermentative products,
256
which are not normally intermediary metabolites for strictly aerobic bacteria. Thus, the presence
257
of these organic acids serves as an indication that Bcc was able to perform fermentation. The
258
technique used for the measurement of low-molecular-weight organic acids in test medium and
259
in vitro ASL was established for anoxic sediment pore-water (47). A gradient liquid
260
chromatograph, combined with an UV/VIS detector and an integrator, was used to identify and
261
quantitate organic acids in pre- and post-infected ASL. The column employed was a 22-cm
262
cartridge with a guard column or polymeric reversed-phase guard cartridge in the sample loop as
263
a concentrator. To measure organic acids, 2-nitrophenylhydrazides (NHPs) of C1 and C5 acids
264
were first produced in aqueous solution through the use of a water-soluble carbodiimide-
265
coupling agent. The test solutions were centrifuged, pyridine buffer added, followed by NHP,
266
then 1-ethyl-3- (3-dimethylaminopropyl) carbodiimide hydrochloride, and finally 40% KOH.
267
The samples were mixed and heated in a heating block at 70oC for 10 min. After centrifugation,
268
the supernatant was pumped through the concentrator column in the sample loop. The
269
derivatized acids adhere strongly because of the cationic quaternary ammonium pairing agents
270
adhering to the stationary phase. After the samples were pumped through the pre-column, 1 ml
271
of deionized water was flushed through the column to remove KOH from the concentrator prior
272
to injection onto the analytical column of the gradient liquid chromatograph. The derivatives
273
were then separated with acetate-buffered ion-pairing solvents and detected by absorption at 230
12
274
or 400 nm. A standard mixture of nine acids (lactate, acetate, formate, propionate, iso-butyrate,
275
butyrate, iso-valerate, valerate, and fumarate; Sigma Chemicals, St. Louis, MO) was run in
276
parallel, which allowed the quantification of the organic acids.
277
Detection of mucin degrading enzyme (“mucinase”) activity. To detect bacterial mucinase
278
activity, we first prepared bacterial cell lysates after growth in Luria Bertani medium overnight
279
at 37oC with shaking. The bacteria were then pelleted and resuspended in TE buffer (10 mM
280
Tris-HCL, 1 mM EDTA, pH 8.0). The cell pellets were lysed by sonication on ice, using six 10-
281
sec bursts at 4 W. Cell debris and unbroken cells were removed by centrifugation at 12,000 rpm
282
for 15 min at 4oC. The supernatants were concentrated and 100-µl volumes dialyzed (Mini
283
Dialysis Unit, Pierce, Rockford, IL) against water. Samples (10x conc.) were then incubated
284
with 1 μg purified bovine cervical mucins for 18h at 37oC. As a positive control, bovine cervical
285
mucins were incubated with trypsin (10 μg). Aliquots were then subjected to SDS/PAGE using a
286
4-12% gradient polyacrylamide gel (BIO-RAD). The samples were electrophoresed at 100 V for
287
1h. The resulting gels were stained with periodic acid-Schiff’s Reagent (PAS; Gelcode
288
Glycoprotein Staining Kit, Pierce). In addition, aliquots were reduced and electrophoresed on
289
1% agarose gels for ~3h in TAE buffer (40 mM Tris actetate, 1 mM EDTA, pH 8.0) at a constant
290
80V. After electrophoresis, the gels were washed in SSC buffer (0.6M NaCl, 60 mM Na citrate,
291
pH 7.0) and the resolved molecules were transferred to nitrocellulose in the same buffer for 1.5 h
292
at a negative pressure of 45 mbar using a VacuGene XL vacuum-blotting apparatus as described
293
(48). After washing, the nitrocellulose membranes were stained with PAS or probed with an
294
antibody specific to cervical polymeric mucins (Muc5AC and Muc5B, 48) and developed with
295
horseradish-conjugated secondary antibody (Jackson ImmunoResearch), in conjunction with an
296
enhanced chemiluminesence kit. The developed membranes were digitized using a Lumax
13
297
Powerlook 1000 Scanner and the integrated staining intensities of the bands in the resulting
298
images were determined with MetaMorph image processing software (49).
299
Statistical Analysis. Mean ± standard errors (SEM) or standard deviations (STDEV) were
300
calculated for ionic, amino acid, and glucose concentrations in SMM, non-CF secretions, and in
301
vitro ASL harvested from WD human airway epithelial cultures. Single parameters in CF versus
302
non-CF samples were compared using Student’s t-Test and were considered significantly
303
different when P
1
Localization of Burkholderia cepacia Complex Bacteria in
2
Cystic Fibrosis Lungs and Interactions with Pseudomonas aeruginosa
3
in Hypoxic Mucus
4 5
Ute Schwab1*, Lubna H. Abdullah1, Olivia S. Perlmutt1, Daniel Albert2, C. William Davis1,
6
Roland R. Arnold3, James R. Yankaskas1, Peter Gilligan4, Heiner Neubauer5, Scott H. Randell1**,
7
and Richard C. Boucher1**
8
Cystic Fibrosis/Pulmonary Research and Treatment and Research Center1, Marine Sciences2,
9
School of Dentistry3, Pathology and Laboratory Medicine, Memorial Hospital4 , University of
10
North Carolina, Chapel Hill, USA; Friedrich-Loeffler-Institut, Institute of Bacterial Infections
11
and Zoonoses, Naumburger Str. 96a, D-07743 Jena, Germany5.
12 13
Running title: Burkholderia cepacia complex bacteria in the CF lung
14 15
* Corresponding author, present address:
16
Clinical Sciences-Immunology and Microbiology
17
College of Veterinary Medicine
18
Cornell University
19
Ithaca, NY 14853
20
Phone 607-229-2399
21
Email: [email protected]
22
** Senior authors contributed equally to this work.
23
This work is dedicated to the memory of our friend and colleague Dr. Anthony Paradiso. 1
24
ABSTRACT
25
The localization of Burkholderia cepacia complex (Bcc) bacteria in the cystic fibrosis (CF) lung,
26
alone or during co-infection with Pseudomonas (P.) aeruginosa, is poorly understood. We
27
performed immunohistochemistry for Bcc and P. aeruginosa on 21 co-/or singly-infected CF
28
lungs obtained at transplantation or autopsy. Parallel in vitro experiments examined growth of
29
two Bcc species, Burkholderia cenocepacia and Burkholderia multivorans, in environments
30
similar to those occupied by P. aeruginosa in the CF lung. Bcc was predominantly identified in
31
the CF lung as single cells or small clusters within phagocytes and mucus, but not as “biofilm-
32
like structures”. In contrast, P. aeruginosa was identified in biofilm-like masses, but densities
33
appeared to be reduced during co-infection with Bcc. Based on chemical analyses of CF and
34
non-CF respiratory secretions, a test medium was defined to study Bcc growth and interactions
35
with P. aeruginosa in an environment mimicking the CF lung. When test medium was
36
supplemented under anaerobic conditions with alternative electron acceptors, B. cenocepacia and
37
B. multivorans used fermentation rather than anaerobic respiration to gain energy, consistent
38
with the identification of fermentation products by HPLC. Both Bcc species also expressed
39
mucinases that produced carbon sources from mucins for growth. In the presence of P.
40
aeruginosa in vitro, both Bcc species grew anaerobically, but not aerobically. We propose that
41
Bcc: 1) invades a P. aeruginosa infected CF lung when the airway lumen is anaerobic; 2)
42
inhibits P. aeruginosa biofilm-like growth; and 3) expands the host bacterial niche from mucus
43
to also include macrophages.
44
2
45
INTRODUCTION
46
Individuals with cystic fibrosis (CF) typically exhibit polymicrobial lung infections, with
47
P. aeruginosa becoming highly prevalent as the disease progresses (1). A subset of CF patients
48
become infected with Burkholderia cepacia complex (Bcc) bacteria (2). Bcc infected CF patients
49
exhibit significantly reduced long-term survival (3), reflecting both a more rapid decline in lung
50
function and, in some cases, the “Cepacia syndrome”, characterized by unrelenting pneumonia,
51
sepsis, and death.
52
However, there is a paucity of information on the localization of Bcc alone, or in
53
conjunction with P. aeruginosa, in the CF lung. Based on the observation that both pathogens
54
can be isolated from CF sputum samples, it is generally believed that Bcc and P. aeruginosa co-
55
localize in macro-colonies (“biofilm-like structures”) within the lungs of chronically infected CF
56
patients. This notion has been supported by in vitro studies demonstrating mixed biofilms in
57
vitro (4-6) and bacterial use of the same chemical signals to control biofilm formation and
58
expression of virulence factors (5). It has also been shown that growth inhibitory substances
59
produced by P. aeruginosa can protect their biofilms against invasion by Bcc, but when no
60
inhibitory components are produced, the two species are able to co-exist (7).
61
There is evidence, however, that these organisms may not co-exist in biofilm-like
62
structures within the CF lung. For example, P. aeruginosa has been shown to grow in sessile
63
biofilm matrices, both in vivo and in artificial sputum medium mimicking the CF lung habitat (8-
64
14). In contrast, Bcc biofilm-like structures have not yet been identified within the intraluminal
65
CF mucus inhabited by P. aeruginosa. The absence of mixed biofilms in excised lung
66
specimens is perplexing because Bcc isolates have been shown to form biofilms in vitro,
67
regardless of species (15, 16), and we previously reported the formation of biofilm-like
3
68
structures on the surface of well-differentiated (WD) human airway epithelial cultures (17).
69
Therefore, it is unclear whether Bcc biofilms in vivo have been missed, or that the current in
70
vitro systems that generate biofilms do not adequately mimic the CF lung environment.
71
Other data suggest that P. aeruginosa and Bcc may occupy different niches within the CF
72
lung. Several clinical and laboratory observations suggest that the niche occupied by Bcc in the
73
CF lung is within pulmonary phagocytes and respiratory epithelial cells rather than in
74
intraluminal mucus. These observations include clinical unresponsiveness of Bcc to
75
antimicrobial therapy despite an isolate’s susceptibility in vitro, isolation of serum-sensitive
76
isolates in bacteremic infection (18), and the close taxonomic relationship between Bcc and the
77
intracellular pathogen B. pseudomallei (19). Intracellular survival of Bcc is reported in multiple
78
pulmonary cell types, including a transformed human bronchial epithelial cell line from a CF
79
patient (20), CFTR-positive and CFTR-negative bronchial epithelial cell lines (21), the
80
transformed human bronchial epithelial cell line BEAS-2B (22), polarized lung epithelial cells
81
(23), A549 pulmonary epithelial cells (24, 25), primary type II pneumocytes (26), and pulmonary
82
macrophages (27-30). These data are complemented by a clinical and pathologic study that
83
localized Bcc within macrophages in freshly excised CF lungs (31). However, because the
84
observations of Sajjan et al. (31) were restricted to CF lungs infected by the transmissible clonal
85
ET 12 strain of B. cenocepacia (Edinburgh-Toronto) lineage (genomovar III, cblA+) (32), it is
86
not known whether the immunohistologic findings represent characteristic features of this
87
species/strain or late stage pulmonary infection with Bcc in general. Importantly, the
88
localization of P. aeruginosa was not investigated in CF lungs from which sputum evidence for
89
P. aeruginosa and Bcc co-infection existed.
90
Consequently, we designed a study to localize both Bcc and P. aeruginosa in CF lungs,
4
91
excised at transplantation or autopsy, infected with one or both organisms. We also designed
92
parallel in vitro experiments to ask whether Bcc could survive in environments similar to those
93
occupied by P. aeruginosa in the CF lung. When mucus plugs and plaques exceed a certain size,
94
and are adjacent to highly metabolic cells, the intraluminal CF mucus can become markedly
95
anaerobic (10). Thus, the anaerobic physiology of P. aeruginosa has been studied extensively
96
(10, 33-37). However, the literature on energy production by Bcc in hypoxic environments is
97
sparse. Consequently, we also performed experiments using airway surface layer (ASL) material
98
harvested from WD airway epithelial cultures as a relevant medium to study how Bcc gains
99
energy under O2-deprived conditions, and the interspecies relationships between Bcc and P.
100
aeruginosa during growth under anaerobic and aerobic conditions.
101 102
MATERIALS AND METHODS
103
CF patients and specimens. Sputum and lung tissue specimens were obtained from 21 CF
104
patients under University of North Carolina (UNC) Institutional Review Board (IRB) approved
105
protocols consistent with all federal and institutional requirements for informed consent and
106
confidentiality. Ten were female and eleven were male (age range 13 to 48 years), 18 lungs
107
were obtained at lung transplantation and 3 at autopsy (Table 1). Tissues were dissected to
108
isolate bronchial and bronchiolar specimens that were fixed in neutral buffered formalin,
109
embedded in paraffin, and sectioned using conventional procedures.
110
Sputum samples were obtained prior to specimen procurement, consistent with routine
111
patient care. Samples obtained closest to the surgical/autopsy dates were analyzed for the
112
presence of Bcc, P. aeruginosa and other bacteria at the University of North Carolina Hospital
113
Clinical Microbiology Laboratory. When possible, the relative bacterial densities were
5
114
determined by a dilution streak plate method. For this semi-quantitative method, a grade of 0,
115
1+, 2+, 3+, or 4+ was assigned depending on the growth seen in each dilution streak on a plate
116
(0=no bacteria and 4+=bacteria present after three dilution streaks).
117
We selected 2-4 tissue blocks from each lung based upon airway size and the presence of
118
mucus within the airway lumen. When possible, two blocks with large airways and 2 blocks
119
with small airways and alveoli were selected (Table S1).
120
Rabbit antisera. Hyperimmune antisera were produced by immunizing rabbits with inactivated
121
whole-cell Bcc or P. aeruginosa antigens and were stored at -20oC until further use. Specificity
122
of the antisera was determined by Western blot against crude bacterial cell lysates, including 5
123
CF P. aeruginosa isolates and 5 Bcc strains (i.e., B. cepacia ATCC 25416, B. cenocepacia, B.
124
vietnamiensis, B. multivorans, and B. dolosa). Bacteria for lysates were grown in TSB, washed in
125
PBS, and adjusted to an optical density at 600 nm of 1 (equal to ~5 × 109 bacteria/ml). Then, 2
126
ml of the whole bacteria were centrifuged for 10 minutes at 5000 X G. The pellet was lysed (125
127
mM TrisHCl pH 7.0, 5.0 mM NaF, 2 mM EDTA, 1% NP-40, 1 mM Na3VO4, 100 μM TPCK,
128
100 μM quercetin, 1 mM PMSF, 1 μg/ml leupeptin, and 1 μg/ml pepstatin, 2% SDS, 20%
129
Glycerol), boiled for 5 minutes at 100°C, sonicated for 15 seconds, and aliquots were stored at -
130
20°C. For Western blots, 10 µl per lane of bacterial lysates were run on NuPage Bis-Tris gels
131
(Invitrogen) and electrophoretically transferred to a polyvinylidene fluoride membrane. The
132
membrane was blocked with 5% powdered milk in Tris-buffered saline with 0.1% Tween-20
133
(TBST) for 1 hour. Blots were incubated with antisera at 1:50,000 and 1:100,000 dilutions in
134
blocking solution overnight at 4°C, washed with TBST, and incubated for 1 hour with
135
peroxidase goat-anti-rabbit secondary antibody diluted 1:100,000 in blocking solution. After
6
136
washing with TBST, signal was detected with Immobilon Western Detection Reagent (Millipore)
137
and exposure to film. A duplicate gel was silver stained to evaluate loading.
138
Immunohistochemistry. Bcc and P. aeruginosa were localized in formalin fixed paraffin
139
sections of CF lung tissue obtained during transplantation or autopsy (Table 1 and 2). In almost
140
all cases, 4 tissue blocks were selected from each patient, 2 containing larger bronchi with
141
cartilage in the airway wall and 2 containing bronchioles, in which cartilage was absent and
142
alveoli were present. Sections were deparaffinized, rehydrated, and subjected to antigen retrieval
143
by boiling in 0.05 M Na citrate pH 6.0. Endogenous peroxidase was inhibited by incubation in
144
3% H2O2 in methanol, sections permeabilized with 0.1% Triton X-100 (in PBS), and washed in
145
PBS, 0.05% Tween 20 (PBST). After blocking in 5% donkey serum, 1% fish gelatin, 1% BSA
146
in PBST, sections were incubated overnight at 4oC with 1:3000 and 1:6000 dilutions of P.
147
aeruginosa or Bcc antisera, respectively. Optimal dilutions for each primary antibody were
148
determined using positive control sections containing easily identifiable Pseudomonas colonies
149
(from a CF patient infected only with P. aeruginosa; Table 1, patient #21) or numerous intra-
150
and extra-cellular B. cenocepacia bacteria (autopsy tissue from a Cepacia syndrome CF patient,
151
Table 1, patient #1). These positive control sections were included in each staining run.
152
Negative controls consisted of double the concentration of normal rabbit serum in place of the
153
primary antibody on replicate sections of every tissue block. After washing with diluent (1:3
154
blocking solution in PBST), sections were incubated with biotinylated Fab fragment donkey anti-
155
rabbit secondary antibody (Jackson ImmunoResearch, West Grove, PA), washed in diluent, and
156
incubated with streptavidin horseradish peroxidase (Jackson ImmunoResearch). Slides were
157
washed with PBST, rinsed in 0.05 M Tris-HCl buffer, pH 7.6 and incubated with
158
diaminobenzidine tetrahydrochloride (10 mg/50 ml) in the same buffer with 0.006% H2O2. After
7
159
rinsing in water, sections were counterstained with ethyl green, dehydrated in ethanol, and
160
coverslipped in Permount. A Nikon microphot SA microscope connected to a 3CC-Chilled
161
Camera (Sony, Marietta, GA) interfaced to a computer was used to capture light microscopic
162
images via Adobe Photoshop.
163
Slides were evaluated by two independent observers and were graded as strong (+++),
164
moderate (++), weak (+) staining for P. aeruginosa and/or Bcc in endobronchial mucus and
165
other locations. Each slide was examined for bacterial appearance (single rods, clusters, or
166
macrocolonies), location of bacteria (inside inflammatory, bronchial epithelial or alveolar cells)
167
and morphology of airway epithelium (intact, partial or total denudation).
168
Supernatants of mucopurulent material (SMM) and airway secretions obtained from CF
169
and non-CF patients. Under UNC IRB approved protocols, mucopurulent respiratory secretions
170
were harvested from CF lungs excised during transplantation (female, n=10; male, n=10; average
171
age, 29.2 years), and from donor lungs that were unused for transplantation (non-CF control;
172
female, n=6; male, n=4; average age, 32.6 years). Approximately 1-5 ml of mucopurulent
173
material (CF) or pooled secretions (control) was directly aspirated from intrapulmonary airways
174
with syringes, ultracentrifuged for 1 h at 100,000 rpm (434,902 x g, Beckman TL-100 Tabletop
175
Ultracentrifuge, TLA 100.2 rotor), and the supernatants were stored at -20oC prior to analyses
176
(38, 39).
177
In vitro Production of Airway Surface Layer (ASL) materials. Because ASL is a relevant
178
culture medium for our studies, we developed techniques to harvest large quantities of ASL from
179
WD airway epithelial cell cultures prepared as previously described (40). Cells were removed
180
from portions of main stem or lobar bronchi from excess donor tissue obtained at the time of
181
lung transplantation or from CF lung tissue, again under UNC IRB approved protocols. Cells
8
182
were grown on 100-mm tissue culture dishes in bronchial epithelial growth medium (BEGM).
183
Passage two cells were seeded on collagen IV coated 30 mm Millicell CM membrane supports
184
(Millipore) at a density of ~106 `cells/cm2 and grown in air-liquid interface (ALI) media in the
185
absence of antibiotics. Upon reaching confluence (~5 days), the culture medium was only
186
replaced in the serosal chamber. At ~21 days, when the apical cell surface of cultures was fully
187
differentiated into ciliated and mucus producing cells (as determined by light microscopy), 200
188
l of Kreb’s Bicarbonate Ringer (KBR) solution was deposited onto the apical cell surface
189
followed by collection of ASL 24 h later with a pipette tip attached to a 3 cc syringe.
190
Preliminary experiments revealed that KBR was conditioned by the epithelium to resemble ASL
191
in vivo with respect to ionic composition within 6 to 12 h (41). After ASL collection, the apical
192
cell surface of cultures was washed with endotoxin free PBS and, after 24 hours, KBR was added
193
for a second harvest of ASL. The harvested ASL was stored at 4oC.
194
Chemical analysis of SMM and in vitro produced ASL. The ions Na+, K+, Cl-, and Mg2+ were
195
measured using the Vitros950/950AT Chemistry System by standard protocols. The electron
196
acceptors nitrate and sulfate were measured in secretions and in vitro ASL by ion
197
chromatography using a Dionex 2010i Ion Chromatograph. Test samples and standards were
198
diluted 1:50 in sodium bicarbonate/sodium carbonate eluent, filtered (0.45 µm filters) to remove
199
particulates, and 1 ml of sample injected onto the columns. Standards were run to determine
200
NO3- and SO42- peaks, sample areas then compared to standards, a standard curve graphed, and
201
the sample mass calculated according to the slope. Iron concentrations were measured
202
spectrophotometrically using a commercial kit (MPR Iron without deproteinisation, Roche
203
Diagnostics Corp., Indianapolis). Amino acids were measured by HPLC (Millipore, Waters 7R
204
WISP). Osmolality was measured by using a Micro osmometer Model 3300 (Advanced
9
205
Instruments, Inc.). The data obtained from in vivo secretions and ASL measurements were used
206
to develop an artificial nutrient limited medium that was used to study bacterial growth in a
207
defined, pathophysiologically-relevant microbiology system. Thus, the range of growth rates
208
produced are predicted to be much slower than those obtained using nutrient-rich laboratory
209
media (e.g. Luria Bertani medium) and may provide a more accurate indication of Bcc bacterial
210
physiology relevant to the CF lung. P. aeruginosa growth and biofilm formation have already
211
been studied in synthetic media that nutritionally mimic CF sputum (11, 12, 42).
212
Purification of mucins. Commercially available mucins are poorly defined chemically and non-
213
gel forming. MUC5AC and MUC5B are the major gel-forming mucins in both bovine cervical
214
mucus and human respiratory mucus (43-45). To produce large volumes of purified
215
Muc5AC/5B mucins, 50 mL of fresh bovine cervical mucus was solubilized in 6 M guanidinium
216
chloide and mucins were purified by double isopycnic density gradient centrifugation (46).
217
Fractions were collected, weighed for density, and assessed for DNA (A260), protein (A270), and
218
sialic acid. Sialic acid-positive fractions were pooled and dialyzed against deionized water. The
219
second gradient centrifugation yielded a sharp sialic acid peak that was well separated from
220
DNA. A known volume of this fraction was dried and its weight determined on a microbalance
221
(CAHN, C33) to determine the mucin content. Mucin solutions, reconstituted with PBS at
222
concentrations ranging from 0.01 to 1 mg in a volume of 100L, were prepared and used for
223
bacterial growth studies.
224
Bacterial strains. Two Bcc species, B. cenocepacia BC-7 (genomovar III, cblA+ major CF
225
lineage ET12; provided by Dr. Richard Goldstein, Boston University) and B. multivorans J-1
226
(genomovar II; UNC Hospitals) and the P. aeruginosa strains PAO1 (ATCC) and Pae 33 (UNC
227
Hospitals) were used in this study. Pae 33 was from an individual with severe lung disease being
10
228
evaluated for lung transplantation. The patient had two additional mucoid strains of
229
Pseudomonas and this smooth strain was resistant to many antibiotics, suggesting long standing
230
infection. Stocks of the bacteria were kept at –70oC in skim milk. For all studies, bacteria were
231
grown at 37oC on sheep blood agar plates.
232
Inoculation of in vitro produced ASL and “pure” mucin solutions. To characterize the
233
growth of B. cenocepacia BC-7 and B. multivorans J-1 in ASL produced in vitro and in mucin
234
solutions, we incubated s small volume of ASL or mucin solution (30 µl) with a low number,
235
(approximately 200 to 500 cfu/1 μl PBS) of bacteria (grown overnight on sheep blood agar and
236
then suspended/diluted in PBS). The cultures were grown for 3 days and their growth
237
monitored under aerobic and anaerobic (10% H2, 5% CO2, 85% N2, anaerobic chamber from
238
Coy Laboratory Products, Inc., Ann Arbor, MI) conditions at 37oC in round bottom 96 well
239
plates. All growth studies were performed in triplicate. For anaerobic growth experiments, both
240
ASL and bacterial suspensions were equilibrated in the anaerobic chamber before ASL was
241
inoculated with Bcc and/or P. aeruginosa. The number of viable bacteria was determined by
242
serial dilutions plated onto sheep blood agar and cultured under aerobic conditions.
243
To test mixed cultures for growth in ASL under aerobic/anaerobic conditions, equal
244
numbers of P. aeruginosa (approximately 200 cfu/0.5 μl PBS) and Bcc (approximately 200
245
cfu/0.5 μl PBS) as well as a different ratio of Bcc to P. aeruginosa ( 100:1 were co-cultured, and
246
after 72 hours, serial dilutions were prepared and plated in parallel onto trypticase soy agar and
247
Bcc selective agar (provided by the Microbiology/ Immunology Laboratory of UNC Hospitals)
248
for bacterial enumeration.
249
Preparation of bacterially conditioned test medium and in vitro ASL supernatants. In vitro
250
ASL was inoculated with the two Bcc species and P. aeruginosa bacteria as described above in
11
251
quadruplicate, and after 72 h of incubation, the four samples were pooled, filtered (0.2 µm
252
Acrodisk) and ASL filtrates stored at –20oC until use.
253
Measurements of organic acids in infected test medium and in vitro ASL. For these studies,
254
bacteria-conditioned and non-inoculated test medium and ASL were assayed for low-molecular
255
weight organic acids, e.g., acetate, lactate, or formate. These acids are fermentative products,
256
which are not normally intermediary metabolites for strictly aerobic bacteria. Thus, the presence
257
of these organic acids serves as an indication that Bcc was able to perform fermentation. The
258
technique used for the measurement of low-molecular-weight organic acids in test medium and
259
in vitro ASL was established for anoxic sediment pore-water (47). A gradient liquid
260
chromatograph, combined with an UV/VIS detector and an integrator, was used to identify and
261
quantitate organic acids in pre- and post-infected ASL. The column employed was a 22-cm
262
cartridge with a guard column or polymeric reversed-phase guard cartridge in the sample loop as
263
a concentrator. To measure organic acids, 2-nitrophenylhydrazides (NHPs) of C1 and C5 acids
264
were first produced in aqueous solution through the use of a water-soluble carbodiimide-
265
coupling agent. The test solutions were centrifuged, pyridine buffer added, followed by NHP,
266
then 1-ethyl-3- (3-dimethylaminopropyl) carbodiimide hydrochloride, and finally 40% KOH.
267
The samples were mixed and heated in a heating block at 70oC for 10 min. After centrifugation,
268
the supernatant was pumped through the concentrator column in the sample loop. The
269
derivatized acids adhere strongly because of the cationic quaternary ammonium pairing agents
270
adhering to the stationary phase. After the samples were pumped through the pre-column, 1 ml
271
of deionized water was flushed through the column to remove KOH from the concentrator prior
272
to injection onto the analytical column of the gradient liquid chromatograph. The derivatives
273
were then separated with acetate-buffered ion-pairing solvents and detected by absorption at 230
12
274
or 400 nm. A standard mixture of nine acids (lactate, acetate, formate, propionate, iso-butyrate,
275
butyrate, iso-valerate, valerate, and fumarate; Sigma Chemicals, St. Louis, MO) was run in
276
parallel, which allowed the quantification of the organic acids.
277
Detection of mucin degrading enzyme (“mucinase”) activity. To detect bacterial mucinase
278
activity, we first prepared bacterial cell lysates after growth in Luria Bertani medium overnight
279
at 37oC with shaking. The bacteria were then pelleted and resuspended in TE buffer (10 mM
280
Tris-HCL, 1 mM EDTA, pH 8.0). The cell pellets were lysed by sonication on ice, using six 10-
281
sec bursts at 4 W. Cell debris and unbroken cells were removed by centrifugation at 12,000 rpm
282
for 15 min at 4oC. The supernatants were concentrated and 100-µl volumes dialyzed (Mini
283
Dialysis Unit, Pierce, Rockford, IL) against water. Samples (10x conc.) were then incubated
284
with 1 μg purified bovine cervical mucins for 18h at 37oC. As a positive control, bovine cervical
285
mucins were incubated with trypsin (10 μg). Aliquots were then subjected to SDS/PAGE using a
286
4-12% gradient polyacrylamide gel (BIO-RAD). The samples were electrophoresed at 100 V for
287
1h. The resulting gels were stained with periodic acid-Schiff’s Reagent (PAS; Gelcode
288
Glycoprotein Staining Kit, Pierce). In addition, aliquots were reduced and electrophoresed on
289
1% agarose gels for ~3h in TAE buffer (40 mM Tris actetate, 1 mM EDTA, pH 8.0) at a constant
290
80V. After electrophoresis, the gels were washed in SSC buffer (0.6M NaCl, 60 mM Na citrate,
291
pH 7.0) and the resolved molecules were transferred to nitrocellulose in the same buffer for 1.5 h
292
at a negative pressure of 45 mbar using a VacuGene XL vacuum-blotting apparatus as described
293
(48). After washing, the nitrocellulose membranes were stained with PAS or probed with an
294
antibody specific to cervical polymeric mucins (Muc5AC and Muc5B, 48) and developed with
295
horseradish-conjugated secondary antibody (Jackson ImmunoResearch), in conjunction with an
296
enhanced chemiluminesence kit. The developed membranes were digitized using a Lumax
13
297
Powerlook 1000 Scanner and the integrated staining intensities of the bands in the resulting
298
images were determined with MetaMorph image processing software (49).
299
Statistical Analysis. Mean ± standard errors (SEM) or standard deviations (STDEV) were
300
calculated for ionic, amino acid, and glucose concentrations in SMM, non-CF secretions, and in
301
vitro ASL harvested from WD human airway epithelial cultures. Single parameters in CF versus
302
non-CF samples were compared using Student’s t-Test and were considered significantly
303
different when P
