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TITLE: 1
Mycofumigation by the volatile organic compound producing fungus Muscodor albus induces 2
bacterial cell death through DNA damage 3
RUNNING TITLE: 4
Antibacterial modes of action of Muscodor albus 5
6
AUTHORS: 7
Cambria J. Alpha1, Manuel Campos
2, Christine Jacobs-Wagner
2, Scott A. Strobel
1*
8
1Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT 9
06520, USA 10
2Microbial Diversity Institute, Department of Molecular, Cellular and Developmental Biology, 11
and Howard Hughes Medical Institute, Yale University, New Haven, CT 06520, USA 12
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ABSTRACT WORD COUNT: 322 15
TEXT WORD COUNT: 7060 (excluding ref, table footnotes, & figure legends): 16
17
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AEM Accepts, published online ahead of print on 1 December 2014Appl. Environ. Microbiol. doi:10.1128/AEM.03294-14Copyright © 2014, American Society for Microbiology. All Rights Reserved.
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ABSTRACT 19
Muscodor albus belongs to a genus of endophytic fungi that inhibit and kill other fungi, 20
bacteria, and insects through production of a complex mixture of volatile organic compounds 21
(VOCs). This process of mycofumigation has found commercial application for control of 22
human and plant pathogens, but the mechanism of the VOC toxicity is unknown. Here the 23
modes of action of these volatiles was investigated through a series of genetic screens and 24
biochemical assays. A single gene knockout screen revealed high sensitivity for E. coli lacking 25
enzymes in the pathways of DNA repair, DNA metabolic process, and response to stress when 26
exposed to the VOCs of M. albus. Furthermore, the sensitivity of knockouts involved in the 27
repair of specific DNA alkyl adducts suggest that the VOCs may induce alkylation. Evidence of 28
DNA damage suggests that these adducts lead to breaks during DNA replication or transcription 29
if not properly repaired. Additional cytotoxicity profiling indicated that during VOC exposure, E. 30
coli became filamentous and demonstrated an increase in cellular membrane fluidity. The 31
volatile nature of the toxic compounds produced by M. albus and their broad range of inhibition 32
make this fungus an attractive biological agent. Understanding the antimicrobial effects and the 33
VOCs modes of action will inform the utility and safety of potential mycofumigation 34
applications for M. albus. 35
36
IMPORTANCE 37
The volatile organic compounds of Muscodor albus have been shown to inhibit a broad 38
range of human and plant pathogens, including bacteria, fungi, and insects. Mycofumigation by 39
M. albus has been proposed for containment of agricultural pathogens, treatment of bacterial 40
infections, and decontamination of food transports. This work provides an understanding of the 41
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antibacterial effects of the volatile metabolites and will inform the utility and safe application of 42
the fungus. Additionally the insight into volatile antimicrobial properties allows for the 43
development of further mycofumigation treatments in both medicine and agriculture. This study 44
is the first to explore the modes of action of these fungal volatiles. 45
46
INTRODUCTION 47
The endophytic fungus Muscodor albus (CZ-620) inhibits growth of a broad range of 48
pathogenic fungi and bacteria, as well as some nematode and arthropod species (1–6). The 49
inhibition is achieved exclusively through a complex mixture of volatile organic compounds 50
(VOCs) that M. albus secretes into the headspace of the culture. The volatile compounds emitted 51
by M. albus and other closely related organisms in the genus consist of a combination of short-52
chain alcohols, organic acids, esters, ketones, and several aromatic hydrocarbons as monitored 53
by gas chromatography-mass spectrometry (GC-MS) (6). The compounds range from two to 54
nine carbons and include both straight and branched chain varieties. The larger aromatic products 55
are predicted to be sesquiterpenes and derivatives of naphthalene and azulene, but have not been 56
confirmed by comparison to standards. Although many fungal species have been reported to 57
produce VOCs, none have demonstrated the wide-ranging bioactivity as is seen with isolates of 58
M. albus (7, 8). 59
The biological function of the toxic compound production by M. albus is unknown. M. 60
albus was first identified as an endophyte, an organism that lives within the inner tissues of 61
plants (9). Although endophytic interactions between fungi and host plants are typically 62
asymptomatic for part, or all, of their life cycle, these can also be commensal relationships. 63
There are examples where fungi improve the host plant’s growth, fitness, or response to stress 64
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response (10–12). In the case of M. albus, its VOC emissions may serve as a defense mechanism 65
for the host plant against insects or potential pathogens, and these same products have been 66
hypothesized to help M. albus’s survival by preventing colonization of the host plant by 67
organisms competing for the same environmental niche (6). 68
Regardless of the biological purpose, the toxicity of M. albus to other organisms is being 69
harnessed for commercial uses. “Mycofumigation” is a process wherein M. albus VOC 70
production inhibits pathogen growth in agricultural seeds, plants, and soil (3, 5, 13). 71
Mycofumigation has also been applied to fruit storage and transportation, where the presence of 72
M. albus increases shelf-life and alleviates pressure for expedited shipping (2). Agraquest, a 73
company recently acquired by Bayer, is exploring the use of M. albus in numerous agricultural 74
applications including, but not limited to, control of fungal and bacterial pathogens in post-75
harvest and soil diseases, building mold remediation, and seed/grain sanitation (patent 76
application #20120058058). In particular, they seek to use M. albus for soil sterilization, 77
replacing the use of methyl bromide, a pesticide that is highly toxic and detrimental to the ozone 78
layer (EPA 69592-RL/T/I). The volatile nature of the toxic compounds produced by M. albus 79
and their broad range of inhibition make this fungus an attractive biological agent for these 80
diverse applications. 81
The emergent commercial use of this organism necessitates a more complete 82
understanding of the underlying biology of M. albus. It appears that the toxicity of M. albus 83
exposure results from the combined action of more than one of the secreted compounds. None of 84
the individual compounds or classes of compounds alone mimic the toxicity of M. albus (6). 85
However, the individual compounds produced by M. albus do have antimicrobial effects at 86
higher levels. Each class of compound produced by M. albus was evaluated for toxicity using 87
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artificial mixtures. On a comparative basis, the esters had more inhibitory activity than any other 88
group (6). Alcohols, including ethanol and larger alkanols, are toxic due to their role in 89
membrane disruption, which leads to the dissipation of the proton gradient (14, 15). The 90
unidentified, larger molecular weight VOCs are predicted to be polycyclic sesquiterpenes 91
(molecular formula C15H24), naphthalene derivatives, or azulene derivatives. Naphthalene has 92
long been used as a household fumigant. The compound is toxic at high concentrations to both 93
the adult and larval forms of many moths, though bacteria have no growth inhibition at 94
concentrations as high as 50 mg/L (16). Azulene is slightly more toxic to the gram-negative 95
bacteria, Vibrio fischeri, with an IC50 value of 1.5 mg/L (17). It has been proposed that these 96
compounds act synergistically in the context of the mixture produced by the fungus (6). It may 97
be due to this combination of toxic agents that M. albus is lethal to such a broad range of 98
organisms. 99
Here we explore the basis of M. albus VOC toxicity using genetic screens to identify the 100
pathways affected by the VOCs. This is coupled with cytotoxicity profiling to monitor bacterial 101
response to VOC exposure. It appears that the VOC mixture targets diverse cellular pathways. 102
These results explain the modes of action and complement the known broad toxicity of the 103
fungus. This analysis will inform the use of M. albus VOCs for mycofumigation. 104
105
METHODS 106
Fungal culture, Muscodor albus CZ-620, was obtained through Professor Gary Strobel at 107
Montana State University. All fungal cultures were maintained on potato dextrose agar (PDA) 108
plates [24 g potato dextrose broth (EMD), 15 g agar (BD Difco), 1 l distilled water] stored at 109
23 °C. Cultures were propagated by transfer of 5 mm culture plugs derived from these plates. 110
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Strains of Escherichia coli were obtained from the American Type Culture Collection 111
(ATCC) (Rockville, Md.), ATCC 25922 and ATCC 35218. Laboratory and clinical isolates of 112
Acinetobacter, Citrobacter, Enterobacter, Klebsiella, Pseudomonas, and Staphylococcus were 113
generously provided by Dr. Thomas Murray (Quinnipiac University). 114
The library of Escherichia coli K-12 knockout clones in the Keio Collection (18) were 115
purchased from Thermo Scientific. E. coli BW25113 was acquired from the Yale Coli Genetic 116
Stock Center (Yale University), and used as control wild type E. coli strain. 117
Bacterial cultures were maintained on autoclaved Luria broth (LB) [10g Tryptone, 5g 118
Yeast Extract, 10 g NaCl, 15 g agar, 1 l distilled water] plates grown at 37 °C and stored at 4°C. 119
When necessary LB was supplemented with Ampicillin 100 μg/ml or Kanamycin 25 μg/ml. For 120
longer storage, 750 μl of overnight culture grown in LB was mixed with 250 μl 80% glycerol, 121
flash frozen in liquid nitrogen and stored at -80°C. 122
Restriction enzymes and DNA ligase were purchased from New England Biolabs 123
(Ipswich, MA, USA). Plasmid pHC79 was obtained from ATCC (Manassas, VA). Plasmid 124
pUC19 was obtained from New England Biolabs. Ampicillin and kanamycin were obtained 125
through Sigma Chemical Company (St. Louis, Mo.). Sytox Green stain is commercially 126
available through Life Technologies. 127
Growth inhibition assays 128
CFU Survival 129
The survival of each bacterial variant was measured after exposure to M. albus VOCs by 130
assaying colony forming units (CFU) (19). M. albus was inoculated on a 10-ml PDA slant in a 131
20-ml glass GCMS vial fused to an identical 20-ml glass GCMS vial (split vial system) and 132
grown at 23 °C for three days. Overnight cultures of test bacteria, grown with constant agitation 133
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of 150 rpm at 23 °C, were diluted in fresh LB medium to OD 0.05. Diluted cultures were 134
exposed to three-day-old M. albus headspace within the split glass vial system. Diluted cultures 135
were placed within the split glass vial system, opposite the 10-ml slant of PDA with day 3 M. 136
albus. This split vial was maintained at constant agitation of 150 rpm at 23 °C with no physical 137
contact between the two cultures, only open headspace. 150 μl aliquots of bacteria culture were 138
taken at intervals, and serial dilutions were plated onto LB agar plates. Samples were removed 139
by syringe through rubber septum as to minimize VOC dispersal from the system. The colony 140
forming units (CFUs) of the post-exposure bacteria culture were quantified after growth at 37 °C 141
overnight. The ratio of CFU after exposure to M. albus VOCs to the CFUs when M. albus is 142
absent was used to determine percent “survival.” 143
For CFU survival of E. coli 25922 in stationary phase the same process as described 144
above was executed with the following exceptions. Overnight cultures of bacteria were not 145
diluted, but rather directly transferred to the split vial system for VOC exposure. 146
Kinetic Growth Profile 147
The growth of bacterial variants was measured using BioTek Synergy 4 Hybrid 148
Microplate reader (Winooski, VT). 5 mm plugs of M. albus were inoculated on 5-ml PDA slants 149
in the surrounding wells of a 12-well plate (Costar 3513 Corning,Ny). Plates were incubated at 150
23 °C for three days at which bacteria were inoculated in the internal wells and exposed to the 151
VOCS. Overnight cultures of bacteria, grown at 23 °C, were diluted to OD 0.05 and 2 ml of the 152
diluted culture were added to the internal wells. Plates were sealed using Petri-Seal adhesive 153
sealing film (Sigma-Aldrich) and incubated with constant agitation of 150 rpm at 23 °C. Their 154
growth was recorded spectrophotometrically at 600 nm every three minutes for the incubation 155
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time of 30 hours. Bacterial growth with non-inoculated PDA slants in the surrounding wells were 156
used as controls in this study. 157
Genomic library screen 158
Isolation and Digestion of gDNA from ATCC E. coli 35218 159
E. coli agarose plugs were prepared as described by Bio-Rad (US/ED Bulletin 1753 160
(California)) with modifications. Overnight cultures grown at 23°C were diluted to OD 0.05 and 161
treated with M. albus VOCs at constant agitation of 150 rpm for 8 hours. At designated time 162
5x108 cells/ml were washed at 4 °C with resuspension buffer (10 mM Tris HCl pH 7.2, 20 mM 163
NaCl, 50 mM EDTA pH 8.0) and re-suspended to half the volume of plugs desired. Cultures 164
were equilibrated to 50 °C. 1xTBE 2% LMP agarose (Lonza. Allendale, NJ) was solubilized by 165
heating and equilibrated to 50 °C (final plug solution was 1%). After equilibration of both 166
solutions to 50 °C, equal amounts of 2% LMP agarose and cell suspension were mixed by gentle 167
agitation. 100 μl was pipetted into each plug mold and set at 23 °C for 30 minutes to solidify. 168
Plugs were then incubated at 37 °C overnight in lysozyme buffer (10 mM Tris HCl pH7.2, 50 169
mM NaCl, 0.2% sodium deoxycholate, 0.5% sodium lauryl sarcosine). Overnight buffer was 170
then removed and samples were placed in Proteinase K Solution (100 mM EDTA pH 8.0, 0.2% 171
sodium deoxycholate, 1.0% sodium lauryl sarcosine, 1 mg/ml proteinase K) at 50 °C for at least 172
4 hours or overnight. 173
Genomic DNA was digested while in the plugs to minimize shearing of DNA and 174
confirmed by PFGE. Plugs were washed with 1x digestion buffer (NEB) and the restriction 175
enzyme was added, PstI or BamH1 (30-50 U per 100 μl plug), for 5 hours at 37 °C. Following 176
digestion plugs were washed three times with 50 mM NaCl, and digestion was confirmed by 177
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PFGE. GELase buffer (Epicentre, Biotechnologies) was added (2 μl per 100 μl plug) and 178
solution was melted at 68 °C. 179
After cooling to 45 °C, GELase enzyme was added and mixed by gentle agitation (1 U 180
per 300 μl melted solution) (Epicentre, Biotechnologies). This mixture was incubated at 45 °C 181
for over 1 hour. Samples were then extracted using phenol:chloroform and the digested DNA 182
solution was precipitated in ethanol. Digested DNA was inserted into the Pst1 or BamH1 sites of 183
vector pHC79 (ATCC® 37030). 184
Packaging of E. coli 35218 library for amplification and screening within Electromax 185
DH10B 186
Packing of E. coli 35218 library in pHC79 was performed as described in the instruction 187
manual for Gigapack III Gold Packaging Extract (Agilent Technologies) with few modifications. 188
Cosmid pHC79 and E. coli DNA library was packaged in phage as described and a 1:50 dilution 189
was mixed with prepared host strain E. coli VCS257. The packaging reaction was titered to 190
ensure 20x coverage of library, plated on LB agar plates supplemented with ampicillin, and 191
viable colonies harvested. Library was isolated using Low-copy Cosmid protocol of the Qiagen 192
Maxi kit (Qiagen, Limburg). 193
Electroporation of library into ElectroMAX DH10b cells (Invitrogen™) was performed 194
per Invitrogen™ instructions. Transformants were grown over night at 37 °C at serial dilutions to 195
maintain single colonies. Overnight plates were then replica plated onto LB agar plates, lids 196
removed and inverted onto three day grown M. albus. Inverted plates were sealed and incubated 197
at 23 °C overnight. Viable colonies were streaked upon LB agar places supplemented with 198
ampicillin and grown over night at 37 °C. 199
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Cosmids from viable transformants were isolated using Qiagen Maxi kit with low-copy 200
cosmid isolation protocol. Primer walking was performed and genomic sequence identified 201
(Genewiz). Identified genes were isolated using PCR from the genomic fragment and inserted 202
into pUC19. Individual genes were transformed in ElectroMAX DH10B cells and screened in the 203
same manner as the library. 204
Independent expression of RecA in E. coli DH10B 205
Using primers pUC19RecaFW and pUC19RecaFW, the recA gene was cloned from E. 206
coli 35218 genomic DNA and inserted into high copy plasmid pUC19. ElectroMAX DH10b 207
cells were electroporated with the pUC19 recA construct as described above. Transformants 208
were then grown against M. albus as described in platereader assay above. 209
Knockout library screen 210
Clones from the Keio collection were provided in 96-well microtiter plates and stored at 211
-80 °C. For propagation, all knockout strains were grown in LB containing 25 μg/ml kanamycin 212
at 37 °C. When grown against fungal samples, knockout strains were growth at 23 °C on LB agar 213
containing kanamycin as described above. 214
Screening was performed using 5x5 mm2 culture plug of M. albus inoculated on a one-215
well PDA plate of identical size to the 96-well microtiter plate (Costar 3593 [Corning, NY]). 216
The fungal sample was grown at 23 °C for three days prior to bacterial exposure. 217
For screening of knock out strains, each 96-well microtiter plate was temporarily 218
removed from the -80 °C storage. Without thawing, a 96-pin replicator was used to inoculate a 219
96-well microtiter plate (Costar 3593 [Corning, NY]) containing 200 ul LB and kanamycin per 220
well. Stock plates immediately returned to -80 °C. Inoculated plates were then put at 37 °C for 221
static growth over night. At same time of plate inoculation, cultures of JW1314 (ΔrecT) and 222
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JW2669 (ΔrecA) were each inoculated in 5 ml LB kanamycin, for controls. These samples were 223
grown at 37 °C with constant agitation overnight. 224
The 96-well plate overnight culture was then pinned on LB agar supplemented with 225
kanamycin. In addition, 2 μl of cultures JW1314 (ΔrecT, known VOC-resistant knockout) and 226
JW 2669 (ΔrecA, known VOC-sensitive knockout) were spotted on same plate to serve as 227
controls. Lids from both this inoculated LB kanamycin plate and the three-day-old culture of M. 228
albus were removed, and cultured plates were inverted over each other and sealed using Petri-229
Seal adhesive sealing film (Sigma-Aldrich). Cultures were grown overnight at 23 °C. 230
Cultures were analyzed for growth after 24 hours exposure to M. albus by visual 231
observation for degree of culture spot growth. Each plate was screened against M. albus three 232
times to ensure reproducibility of the results. Plates in which JW2669 (ΔrecA) was not inhibited 233
by M. albus or JW1314 (ΔrecT) was inhibited were disregarded from the analysis. 234
Gene Ontology analysis 235
We used ontologies from Gene Ontology (20) 236
(http://www.geneontology.org/ontology/gene_ontology.obo, version 2014-04-30), while 237
annotations were obtained from EcoCyc for Escherichia coli strain MG1655 (21). Analysis was 238
performed using custom build algorithm in MATLAB including a hypergeometric test to 239
compute p-values which were subsequently adjusted with the Benjamini-Hochberg-Yekutieli 240
False Discovery Rate procedure (22, 23). MATLAB codes are available upon request. 241
Microscopy 242
E.coli cells were grown for more than 5 generations, up to exponential phase 243
(OD600nm~0.1) at 25 °C with constant agitation. The cultures were then exposed to the fungus M. 244
albus for the specified amount of time and spotted on agarose pads before imaging at room 245
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temperature with an inverted Eclipse80i microscope (Nikon, Tokyo Japan) equipped with a 246
Hamamatsu Orca-ER camera and phase-contrast objective Plan Apochromat 100×/1.40 NA. 247
Images were acquired using the MetaMorph software from Molecular Devices (Sunnyvale, CA 248
USA). DAPI labeling was performed 30 minutes prior to imaging (1 g/mL). Cellular length and 249
DAPI intensity were measured using MicrobeTracker (24). 250
NPN-Uptake Permeability Assay 251
Cell permeability using the dye 1-N-phenyl-naphthylamine (NPN) (Sigma-Aldrich) was 252
measured as previously described (25). E. coli was grown for more than 5 generations, up to 253
exponential phase (OD600nm ~0.1) in LB with appropriate antibiotic at 23 °C. Cultures were then 254
treated to the various conditions and grown for another 3 hours, shaking, at 23 °C. After three 255
hours of exposure samples were washed and diluted to OD of ~0.7/0.8. An aliquot of cells at this 256
point was removed and plated for CFU measurement. Polymyxin B was added at ½ minimal 257
inhibitory concentration (MIC) (0.5 ug/ml) and allowed to incubate 5 minutes before 258
measurement. Cell suspensions were analyzed using PTI-814 fluorescent spectrophotometer 259
(New Jersey). Fluorescence was detected with an excitation of 350 nm and an emission of 420 260
nm with split widths of 4 nm. Fluorescence was then normalized by the CFU data for each 261
condition giving relative fluorescence (RFU) of each sample. 262
Pulsed Field Gel Electrophoresis for DNA Damage detection 263
Agarose plugs for pulsed field gel electrophoresis were prepared as described above in 264
“Isolation and Digestion of gDNA from E. coli 35218.” After incubation with Proteinase K 265
solution, samples were incubated in 1x TBE to prepare for electrophoresis. Samples were run on 266
a 1% agarose (Pulse Field Certified Agarose, Bio-Rad) gel for 7 hours at 14 °C, 6 volts, initial to 267
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final switch of 1-6 volts. Gel was stained using 10,000 fold dilution of SYBR Gold nucleic acid 268
stain (Life technologies) for 40 minutes and imaged using Chemidoc MP (Bio-Rad). 269
Gel Electrophoresis for analysis of DNA nicks and strand breaks 270
Gel Electrophoresis was performed on plasmid pBR322 (New England Biolabs Inc) as 271
previously described (Colis 2014). Supercoiled pBR322 was independently treated with Nt.Bsm1 272
nicking enzyme and restriction enzyme BamH1 (New England Biolabs Inc). Each sample (0.5 273
µg) was loaded on 1.2% agarose gel and run at 6 V/cm. Gel was then stained using SYBR® Gold 274
(Life Technologies) at 1:10,000 dilution. 275
276
RESULTS: 277
Identifying sensitive and resistant bacterial strains 278
We set out to identify the modes of action (MOA) of M. albus VOC toxicity by screening 279
a collection of Escherichia coli isolates (Thomas Murray (Yale University), ATCC) for viability 280
after treatment with M. albus VOCs. The collection consisted of seven E. coli strains, derived 281
from both E. coli B and E. coli K-12, including clinical lab isolates and cultured isolates (Table 282
1). Using the simple bioassay system devised to examine only volatile agents for microbial 283
inhibition (6), test E. coli cultures were monitored for growth during exposure to the M. albus 284
VOCs. No visible growth during exposure was observed for five of the seven strains tested. After 285
24 hours, the VOCs were removed and the absence of further growth revealed that the inhibition 286
was bactericidal to these five strains (Table 1). Of these sensitive strains, we selected the strains 287
E. coli ATCC®
25922™
and DH10B for further analysis. E. coli 25922 is a well-characterized 288
strain that is a control Gram-negative bacterium widely used for various laboratory experiments, 289
especially for antibiotic susceptibility assays (26). The strain E. coli DH10B was selected 290
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because it is efficient for DNA cloning. Two of the test bacteria, E. coli ATCC®
35218™
and K-291
12 E. coli BW25113, remained viable after 24 hours of VOC exposure despite an initial static 292
growth inhibition. E. coli 35218 is also a commonly used ATCC strain for antimicrobial 293
susceptibility studies, and BW25113 is the parent strain for the Keio Collection of single gene 294
knockouts (18). 295
Quantitation of M. albus VOC inhibition was accomplished by measuring colony-296
forming units (CFU) for each strain during exposure to the VOCs. E. coli 25922 was completely 297
susceptible to the VOC exposure and steadily lost viability during treatment (Figure 1). Tolerant 298
strain E. coli BW25113 exhibited static inhibition of growth upon VOC exposure; however, 299
following the initial hours of exposure E. coli BW25113 resumed exponential growth. Similar to 300
BW25113, E. coli 35218 was viable after 24 hours of M. albus VOC exposure. In 35218, 301
however, the initial growth inhibition was much more severe as the culture viability steadily 302
decreased over the first few hours of VOC treatment, then growth recovered and mimicked that 303
of untreated cultures (Supplementary Figure 1). 304
305
Identifying the modes of action 306
The dramatic differences in growth among the different E. coli strains suggested that a 307
genetic variation between the strains was responsible. We set out to use these differences in VOC 308
susceptibility for a series of genetic screens to identify the genes that play a role in E. coli 309
resistance. 310
We first determined if a specific genetic element was sufficient to render a bacterial strain 311
tolerant to M. albus VOCs. We constructed a plasmid-based library of the genomic DNA from 312
the tolerant E. coli strain 35218 and screened for resistance in the highly sensitive E. coli strain 313
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DH10B. We cloned fragments of E. coli 35218’s genome in DH10B and screened the 314
transformants for viability after exposure to M. albus VOCs. The screen resulted in four viable 315
clones out of the approximately 3000 tested. We extracted the library-plasmid from each of the 316
four viable clones and sequenced the genomic library fragments to identify the individual genes 317
encoded in each fragment. The gene encoding the DNA repair protein recombinase A (RecA) 318
was present in all four of the isolated clones. An independent construct containing solely the 319
recA gene was sufficient to rescue viability of E. coli DH10B after VOC exposure for 24 hours. 320
This transformant initially demonstrated static growth during exposure to VOCs, however after a 321
single doubling time it resumed exponential growth (Figure 2). No other gene isolated from the 322
clones was sufficient to induce resistance in DH10B. Isolation of a single repair gene 323
demonstrated that DNA repair is an essential process for viability against M. albus. In the 324
genetic background of DH10B, RecA is sufficient for viability. DH10B is a highly efficient 325
competent cell line that contains the recA1 allele, which has a single point mutation that renders 326
the strain defective in all known in vivo functions of the recA gene. This increases its utility as a 327
cloning strain as it reduces the occurrence of unwanted recombination of cloned DNA (27). 328
However, the highly sensitive strain E. coli 25922, in addition to other clinical and lab isolates 329
(Table 1), contain functioning recA genes and functional DNA repair pathways and yet are still 330
sensitive to M. albus VOCs. This result led us to suspect that while DNA repair is required for 331
resistance, it is not the only component necessary for tolerance to the VOC mixture. 332
Given that a single gene could confer resistance to the M. albus VOCs, we sought to 333
identify all the genes that contribute to tolerance in E. coli. We screened the Keio collection of 334
single gene knockout strains in the background of the tolerant E. coli BW25113. The inhibitory 335
effect of VOC exposure on the single-gene knockouts was scored as follows: complete inhibition, 336
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slight inhibition, or no effect relative to the unexposed controls. Out of the 3869 known-ORF 337
knockouts, a total of 141 gene knockouts had increased sensitivity to VOC exposure. Twenty-338
seven of the single gene knockouts were completely inhibited by M. albus VOCs. All the 339
knockouts that demonstrated increased sensitivity to M. albus VOCs were categorized using 340
Gene Ontology (GO) (20). Using these GO classifications, we performed a gene-set enrichment 341
analysis. This classified the mutants into a total of 17 GO categories that were enriched among 342
knockouts sensitive to M. albus VOCs (Figure 3). The most enriched categories were DNA 343
repair, DNA metabolic process, and response to stress. Additional categories, such as response to 344
DNA damage, SOS response, macromolecule metabolic processes, and DNA recombination, 345
were also prominent in the sensitive set. The enrichment analysis supported our initial 346
observations that DNA repair was necessary for VOC tolerance, but it also highlighted other 347
pathways not previously considered. 348
Sensitive knockouts include the DNA repair genes recA, B, C, D, G, F, J, N, O, and R 349
and sixteen additional genes involved in DNA repair, including uvrD and ruvA. Approximately 350
40% of the genes involved in DNA repair increased the sensitivity to M. albus VOCs when 351
absent. Among these were genes involved in homologous recombination (HR), nucleotide 352
excision repair (NER), base excision repair (BER), and mismatch repair (MMR), each of the 353
DNA repair categories in bacteria. The second most enriched category involved genes that 354
respond to cellular stress, including many envelope stress proteins, outer membrane transporters, 355
proteins that respond to oxidative reagents, and proteins involved in the response to extracellular 356
stimuli including ethanol and osmotic shock. The knockouts of SOS response genes, including 357
ruvA, ruvV, dinB, recA, recN, symE, recF, uvrA, uvrD, and uvrC, all demonstrated increased 358
sensitivity to VOCs. Eleven of the 141 sensitive knockouts were involved in protein and RNA 359
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degradation, including the Lon and other proteases, ClpX, ElaD, HyaD, and GlgG. Unexpectedly, 360
the gene knockouts of four cell division proteins (MinC, Ttk, YcbG, ZapA) and six proteins 361
involved in chemotaxis (CheY, FlgC, FliJ, Flk, YraI, YcbU) also demonstrated increased 362
sensitivity to the M. albus VOCs. Our results seem to indicate a large role for cellular stress 363
responses and DNA repair in M. albus tolerance, however the diversity of other genes suggests a 364
complex MOA with multiple targeted pathways. 365
366
M. albus VOCs induce DNA damage 367
We sought to verify the direct role of M. albus VOCs on these target pathways. We 368
monitored tolerant and sensitive bacterial strains exposed to M. albus VOCs for phenotypic 369
changes in cellular morphology, membrane fitness, and DNA integrity. 370
Live-cell imaging showed that after a 3-hour exposure, the VOCs produced by the fungus 371
resulted in filamentation of the sensitive E. coli 25922 cells (Figure 4A, B) and, to a lesser extent, 372
of the tolerant E. coli 35218 cells (Supplementary Figure 2A, B). With longer treatment, the 373
morphology of the tolerant E. coli 35218 strain was similar to that of non-exposed cells (data not 374
shown). A filamentous phenotype in E. coli could be due to a direct effect on the division 375
machinery or to an indirect inhibition of cell division. When cell division is directly inhibited 376
with the -lactam antibiotic cephalexin, DNA replication and segregation occurs normally, 377
resulting in multiple nucleoids evenly distributed along the cell filament (data not shown and 378
(28)). In contrast, after DAPI staining, we observed that the DNA of the sensitive cells exposed 379
to M. albus remained confined, forming one centrally-located nucleoid flanked by large DNA-380
free regions (Figure 4A and Supplementary Figure 2A). In addition, the DNA amount per cell, as 381
measured by the DAPI signal intensity, was barely higher in exposed cells compare to VOCs-382
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untreated cells, despite a large difference in cell size (Figure 4B, C). As a consequence, the DNA 383
concentration per cell was considerably smaller in cells exposed to VOCs (Figure 4D). This 384
difference was much less noticeable for the tolerant strain (Supplementary Figure 2C, D). These 385
results are consistent with the idea that VOCs induce DNA damage and trigger related stress 386
responses that result in inhibition of DNA replication and cell division. 387
Antimicrobial agents whose MOA is the inhibition of DNA replication induce death 388
through the accumulation of DNA breaks that are either too abundant to be repaired or cannot be 389
repaired due to inactivated replication or repair machinery. We monitored the formation of DNA 390
breaks using pulsed-field gel electrophoresis (PFGE) where the distribution of randomly broken 391
DNA molecules can be determined by DNA migration from the high-molecular weight 392
chromosomal DNA band (29, 30). Exponentially growing cultures were exposed to M. albus 393
VOCs and then embedded in low melting point agarose gel in order to prevent the indirect 394
breakage of genomic DNA during the subsequent cell lysis. After analysis by PFGE, the 395
genomic DNA from healthy non-exposed cells was observed as a single, clear band with 396
negligible evidence of DNA fragmentation (Figure 5a, lanes 2-4). By contrast, the genomic DNA 397
isolated from sensitive, exposed strains had significant accumulation of lower-molecular-weight 398
DNA fragments, indicative of double-stranded DNA damage (Figure 5a, lanes 5-7). In sensitive 399
strains, the accumulation of fragments increased with increasing VOC exposure time. This 400
accumulation was not visible in tolerant strain E. coli BW25113 (Fig 5b), where exposed cells 401
showed negligible evidence of DNA damage, similar to healthy non-exposed cells. This visible 402
accumulation of short DNA fragments in the sensitive strain exposed to VOCs confirmed that 403
double-stranded DNA damage is an important element of M. albus toxicity. 404
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There are at least two mechanisms to explain this accumulation of DNA damage. The 405
first is an indirect model in which M. albus VOCs interact with cellular compounds or machinery 406
that in turn cause damage to DNA. The second involves direct damage to the DNA in the form of 407
nicks or breaks by M. albus VOCs. 408
To test if the VOCs are acting by a direct or an indirect mechanism, we exposed purified 409
supercoiled plasmid pBR322 to the M. albus VOCs. The exposed DNA was then analyzed using 410
gel electrophoresis (Figure 6). Mobility of a supercoiled plasmid through high-concentration 411
agarose does not reflect its linear size due to its tightly wound conformation. A single nick in the 412
plasmid relieves the superhelical tension and causes the DNA to migrate slower in the gel. When 413
completely linearized the plasmid runs according to its true length relative to a double-stranded 414
DNA size ladder. No visible nicking or cleavage was observed upon VOC treatment of purified 415
plasmid after 24 and 48 hours. These data suggest that the DNA damage within sensitive 416
organisms is not due to direct lesions caused by M. albus VOCs. Instead, the damage results 417
from an indirect process. 418
These data, in addition to the observation of DNA replication arrest, suggest that the M. 419
albus VOCs have a toxic effect on bacteria that are actively growing, replicating, and dividing. 420
To test this, we exposed a culture of E. coli 25922 in stationary phase to the VOCs and 421
monitored viable cell counts over time. After seven hours of exposure, approximately 95% of the 422
stationary phase culture was killed compared to more than 99.99% of cells in an exponentially 423
growing culture (Figure 7). The metabolically reduced culture was at least one thousand-fold less 424
sensitive to the VOCs than the exponentially active culture. These data suggest that M. albus’s 425
VOCs are acting in a manner that is dependent on the metabolic activity of the bacteria. 426
427
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M. albus VOCs increase membrane permeability 428
In addition to DNA processes, cell envelope fitness is another common target for 429
antibiotics, including permeation of the cell membrane by alcohols, such as those found in the M. 430
albus VOC mixture. Of the 27 gene knockouts belonging to the cellular response to stress 431
category, 15 were specifically related to membrane functions. To test for effects on membrane 432
integrity, we probed for damage of the bacterial outer membrane after exposure to M. albus 433
VOCs using the cell impermeable dye 1-N-phenylnaphthylamine (NPN). The intensity of the 434
fluorescent probe is correlated to the degree of outer membrane damage and intercalation of the 435
dye into the membrane. We found that untreated healthy cells display a background level of 436
fluorescence due to non-specific surface binding of the dye (Figure 8). The fluorescence 437
increased 3-fold upon treatment with Polymyxin B (at 1/2 MIC), an antibiotic known to 438
permeate outer membranes of Gram-negative bacteria. Tolerant strain E. coli BW25113 439
displayed a greater than 10-fold increase in NPN fluorescence upon exposure to M. albus VOCs 440
for three hours. Similarly, the sensitive recA knockout of BW25113 demonstrated an increase in 441
permeability upon VOC exposure. These results indicate that M. albus VOCs cause an increase 442
in membrane permeability, independent of strain resistance or ability to repair DNA. 443
444
Discussion: M. albus displays multi-target toxicity 445
M. albus produces a mixture of VOCs that are lethal, not only to a broad range of bacteria 446
and fungi, but also some nematode and arthropod species. This antimicrobial activity stems from 447
one or a combination of the VOCs produced by the fungus, which includes alcohols, acids, esters, 448
ketones, and aromatic hydrocarbons (6). The modes of action of these VOCs and the pathways 449
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they target to inhibit bacterial growth was unknown and has been challenging to characterize due 450
to the volatility and complex mixture of the compounds. 451
How does M. albus inhibit such a diverse group of organisms? Previous studies suggested 452
that M. albus’s toxicity is due to the synergistic activities of multiple antimicrobial compounds 453
(6). Using a combination of genomic knockout viability, gain-of-function screening, and 454
comparative growth inhibition assays, we have demonstrated that M. albus toxicity is the result 455
of VOCs permeabilizing the cell membrane and inducing disruption of cellular DNA metabolism 456
through DNA damage. Our data support the multiple-component MOA hypothesis proposed by 457
initial studies on M. albus (6), and identifies the particular pathways targeted by the complex 458
VOC mixtures. 459
Increase in membrane permeability 460
The gene ontology category “cellular response to stress” includes outer membrane 461
proteins related to cellular envelope stress and membrane transporters. The outer membrane of 462
Gram-negative bacteria is an efficient barrier to noxious compounds, antibiotics, and 463
hydrophobic compounds. A selective permeation barrier is notably maintained through 464
nonspecific porin channels as well as specific channels such as TonB-dependent siderophore 465
transports (31). Disruption of these channels can lead to increased susceptibility to antimicrobials. 466
In our screen, E. coli BW25113 knockouts of genes encoding nonspecific channels or porins and 467
specific transporters such as TonB and MdtK, were strongly inhibited by VOCs. 468
Membrane stress could also derive from direct interaction of compounds with the lipid 469
bilayer. Compounds such as butanol and ethanol are known cell membrane disrupters as they 470
disorder the physical structure of the membrane (32). M. albus produces short chain alcohols that 471
could have this activity. These short-chain, branched alcohols are known to have toxic effects on 472
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E. coli due to increased permeability when present at high concentrations (15, 33). In our screen, 473
the knockout of 44 genes functioning as or in association with membrane proteins, rendered the 474
E. coli highly sensitive to M. albus VOCs. The involvement of all these membrane-related 475
proteins emphasizes the importance of maintaining a selective barrier against the VOC mixture, 476
and gives insight into part of M. albus‘s toxic MOA on the bacterial outer membrane. 477
As previously mentioned, 27 of the sensitive gene knockouts belonged to the “cellular 478
response to stress” category. Six of these knockouts, fumC, hscB, iscA, iscU, ygfX and sodC are 479
involved in the cellular stress response to oxidative reagents. Reactive oxygen species have 480
recently been implicated in the mechanism of cellular death by bactericidal antibiotics (34, 35). 481
Although the role of ROS in the toxicity of antibiotics has been debated, an increase in the 482
intracellular levels of ROS has been measured upon treatment with bactericidal antibiotics under 483
aerobic conditions (34, 36–40). For example, the antibiotic ampicillin primarily targets the 484
biosynthesis of the peptidoglycan cell wall; however it also induces increased levels of ROS that 485
correspond to initiation of the SOS response, implicating DNA damage (34). Similarly, n-486
butanol increases membrane fluidity, while at the same time induces a large increase in ROS (41). 487
This intracellular increase can lead not only to DNA damage through sugar modifications, but 488
also to inactivation of enzymes and oxidation of amino acids and polyunsaturated fatty acids 489
(42–45). In all cases, this increase in oxidative stress is the cellular response to the extracellular 490
xenobiotics and highlights the importance of mitigating the interactions of ROS and cellular 491
DNA (46). 492
Interruption of DNA metabolism 493
The most prevalent consequence of M. albus‘s VOC exposure is disruption of DNA 494
metabolism. This is evident in the phenotypic changes of VOC-exposed E. coli. The elongation 495
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of cell length with no increase in the amount of DNA per cell is a well-documented effect of 496
DNA replication inhibition (47–49). In E. coli, cell division and DNA replication are closely 497
coordinated. When DNA replication is interrupted, bacteria initiate the SOS response, which 498
causes an immediate arrest of cellular division and leads to the filamentous phenotype (48). This 499
is a clear and distinct phenotypic effect seen for all cells when exposed to M. albus VOCs. 500
Induction of the SOS response involves de-repression of genes involved in DNA repair, DNA 501
mutagenesis, and the inhibition of cell division. The SOS response acts as an inducible DNA 502
repair system to survive high levels of DNA damage. This response plays a major role in VOC 503
tolerance, as the knockouts of ten SOS response proteins were highly sensitive to the M. albus 504
VOCs. The necessity of DNA repair and SOS proteins for tolerance of M. albus VOCs clearly 505
demonstrates a MOA involving a disruption of DNA metabolism. 506
Further supporting the role of VOCs in DNA disruption, a single DNA repair protein, 507
RecA, was the only hit from the functional genetic screen of a tolerant strain. This indicates that 508
in DH10B functional DNA repair systems are necessary for survival under M. albus VOCs. 509
Although expression of recA conferred tolerance to a sensitive strain DH10B, DNA repair is not 510
sufficient to confer resistance in all sensitive strains. For example, E. coli 25922 was completely 511
inhibited by M. albus VOCS, but based on available strain information all DNA repair 512
mechanisms and SOS proteins (including RecA and Lon) are functional. The DNA repair system 513
that is sufficient for one E. coli strain, is not sufficient for all. This conclusion and the 514
enrichment within 17 gene categories support the multiple-component MOA hypothesis and 515
emphasize the large role played by DNA damage. 516
The necessity for DNA repair leads to the question of what is causing this damage. Our 517
data suggests that the M. albus VOCs do not cause direct single or double stranded breaks to 518
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isolated DNA in solution. However, many of the genes from our knockout sensitivity screen 519
suggest that the DNA is being modified. For example the knockout of DNA methyltransferase, 520
Ada, was highly sensitive to the VOCs. Ada is a multifunction protein that protects E. coli from 521
methylation damage (Metz 2007, McCarthy 1985) Riazuddin 1978). It possesses 522
methyltransferase activity and is converted into a potent transcriptional activator initiating 523
transcription of the methylation resistance genes (ada, alkA, alkB, and aidB) upon methylation 524
(McCarthy 1985). Of these four methylation resistance genes, both ada and alkA were sensitive 525
in our screen. AlkA is a 3-methyl-adenine DNA glycosylase (Nakabeppu,1984). AlkA works to 526
remove not only methyl adducts from adenine but also methyl and ethyl derivatives of guanine 527
and thymine (Metz 2007, Nakabeppu,1984). Similarly, the enzyme TAG is a 3-methyl-adenine 528
glycosylase that was sensitive in our screen (Metz 2007). The requirement of these particular 529
genes for VOC resistance strongly suggests the formation of these lesions, that if not properly 530
repaired would form mutation or lead to breaks during replication (Haffner 2011, Petermann 531
2010). Their sensitivities despite the presence of redundant enzymatic functions suggests the 532
VOCs are inducing the formation of alkyl adducts on the DNA. 533
The repair enzymes that were not hits in the screen reinforce the importance of the 534
sensitive repair-enzyme knockouts and provide further insight into the VOC induced DNA 535
damage. For instance the knockout of DNA glycosylase, MutM, did not create sensitivity. MutM 536
is primarily responsible for removing oxidized bases due to free radical induced lesions (Tajiri 537
1995,Mikawa 1998). Similarly the endonucleases Nfo and Nei, which were not required for 538
resistance, are specific for damaged bases due to specific oxidants (Cunningham 1986, Ischenko 539
2001, Jiang 1997). As this insensitivity could be due to crossover and redundancy of function 540
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between enzymes, it reduces but does not completely eliminate the likelihood that oxidized base 541
modifications are formed upon Muscodor VOC exposure. 542
543
Current microbe control for plants, soil, fruit and vegetable transport utilizes fumigants 544
such as methyl bromide, chloropicrin, or phosphine, among others (50). These chemicals are all 545
toxic to humans, often harmful to plants, and methyl bromide causes significant ozone depletion 546
(Scientific assessment of ozone depletion 2006). M. albus VOCs have been suggested as an 547
alternative to these broad-spectrum chemical fumigants. Without in-depth analysis, this use of M. 548
albus is attractive as the antimicrobials are “naturally” produced by the organism. These natural 549
products, however, are DNA-damaging and their effect on human DNA is yet to be determined. 550
Understanding the mechanisms of action of the VOCs will help establish the appropriate 551
applications of M. albus as an industrial mycofumigant. 552
553
ACKNOWLEDGMENTS: 554
This work was supported by the Naval Grant N00244-09-1-0070 from the United States 555
Department of Defense. 556
557
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FIGURE LEGENDS: 700
Figure 1. Growth and Inhibition of E. coli upon treatment with M. albus VOCs. Growth of 701
K-12 E. coli BW25113 (a. (blue)) and of E. coli 25922 (b. (blue)) as monitored by CFU assay, 702
and post-exposure to the M. albus VOCs (red). 703
Figure 2. Growth curve of E. coli DH10B and E. coli DH10B expressing recA during 704
treatment with VOCs (a.) DH10B growth curve (black) as measured spectrophotometrically by 705
OD600 and upon treatment with M. albus VOCs (red). (b.) Growth curve of DH10B expressing 706
pUC19 recA (black) and during treatment with M. albus VOCs (red). Data is representative of at 707
least three biological replicates. 708
Figure 3. Gene Ontology Category Enrichment Analysis Gene Ontology (GO) category 709
classification of gene knockouts demonstrating increased sensitivity to treatment with M. albus 710
VOCs. Enrichment analysis calculated using a 1% cut-off. GO categories are listed on y-axis in 711
descending order of enrichment. Negative log of the p-value for category enrichment are 712
displayed on x-axis. 713
Figure 4. M. albus VOCs affect cellular morphology and DNA integrity in E. coli 25922 714
cells. (A) Phase contrast and fluorescence images of E. coli 25922 cells after a 3-hour period at 715
25°C in the presence (+) or absence (-) of M. albus VOCs. Cultures were DAPI-stained 716
(2mg/mL) for 20 min prior to imaging on 1% agarose pads. (Scale bar = 2 mm). (B) Density 717
probability distribution of cell length in the absence (red line) or presence of M. albus (orange 718
line). The cell length distribution of E. coli K12 is shown in gray for comparison. (C) Density 719
probability distribution of the DAPI fluorescence signal per cell in the absence (red) or presence 720
(orange) of M. albus in arbitrary units (au). (D) Density probability distribution of the DAPI 721
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30
mean intensity per cell in the absence (red) or presence (orange) of M. albus in arbitrary units 722
(au). 723
Figure 5. Pulsed-field gel electrophoresis (PFGE) of genomic DNA from E. coli treated with 724
M. albus VOCs (a.) E. coli 25922 without exposure to M. albus VOCs (lanes 2-4) or exposed to 725
M. albus VOCS (lanes 5-7) for 3, 4, or 5 hours (from left to right) (b.) E. coli BW25113 was 726
treated as described above, (-) without VOC exposure (lanes 9-11), (+) with VOC exposure 727
(lanes 12-14). Cells/plug was normalized by weight of cell pellet. Lanes 1 and 8 show 24kb Max 728
DNA ladders (Fisher). 729
Figure 6. Fig 6. Gel Electrophoresis analysis for single- and double-stranded DNA breaks 730
of plasmid DNA post exposure to M. albus volatile compounds. Plasmid pBR322 (lane 1 & 6) 731
was exposed to M. albus VOCs in TE pH 8.0 for 24 hours (lane 2) and 48 hours (lane 3). 732
pBR322 was cut with restriction enzyme BamH1 (lane 4) and nicking enzyme Nt.Bsm1 (lane 5) 733
for comparison. 734
Figure 7. Assessment of VOC toxicity on E. coli metabolic state. Percent viability of E. coli 735
25922 cultures exposed to M. albus VOCs for 0 hours (black) and 7 hours (gray) as measured by 736
colony forming units (CFUs). Percent viability was calculated by number of CFUs in culture 737
exposed to VOCs compared to the CFUs of control culture with no VOC treatment. 738
Figure 8. NPN-uptake Permeability of the outer membrane of E. coli upon M. albus VOC 739
exposure. Permeability of K-12 E. coli BW25113 (black) and K-12 E. coli BW25113 ΔrecA 740
(gray) in response to treatment with Muscodor VOCs. Antibiotics were administered at 741
concentrations corresponding to 1/2 MIC. Permeability was monitored by fluorescence of 1-N-742
phenylnaphthylamine (NPN) at 420nm. (RFU) Relative fluorescent unit = intensity of 743
fluorescence/cell density (CFU). 744
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0.001
0.01
0.1
1
10
100
% viable CFUs
E. coli growth phase
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