e01520-15.full.pdf

Enzymatic Degradation of Phenazines Can Generate Energy and

Protect Sensitive Organisms from Toxicity

Kyle C. Costa,

Megan Bergkessel,

a,b,c

Scott Saunders,

Jonas Korlach,

Dianne K. Newman

a,b,c

Division of Biology and Biological Engineering

and Division of Geological and Planetary Sciences,

California Institute of Technology, Pasadena, California, USA; Howard

Hughes Medical Institute, Pasadena, California, USA

; Pacific Biosciences, Menlo Park, California, USA

ABSTRACT

Diverse bacteria, including several

Pseudomonas

species, produce a class of redox-active metabolites called

phenazines that impact different cell types in nature and disease. Phenazines can affect microbial communities in both positive

and negative ways, where their presence is correlated with decreased species richness and diversity. However, little is known

about how the concentration of phenazines is modulated

in situ

and what this may mean for the fitness of members of the com-

munity. Through culturing of phenazine-degrading mycobacteria, genome sequencing, comparative genomics, and molecular

analysis, we identified several conserved genes that are important for the degradation of three

Pseudomonas

-derived phenazines:

phenazine-1-carboxylic acid (PCA), phenazine-1-carboxamide (PCN), and pyocyanin (PYO). PCA can be used as the sole carbon

source for growth by these organisms. Deletion of several genes in

Mycobacterium fortuitum

abolishes the degradation pheno-

type, and expression of two genes in a heterologous host confers the ability to degrade PCN and PYO. In cocultures with

phenazine producers, phenazine degraders alter the abundance of different phenazine types. Not only does degradation support

mycobacterial catabolism, but also it provides protection to bacteria that would otherwise be inhibited by the toxicity of PYO.

Collectively, these results serve as a reminder that microbial metabolites can be actively modified and degraded and that these

turnover processes must be considered when the fate and impact of such compounds in any environment are being assessed.

IMPORTANCE

Phenazine production by

Pseudomonas

spp. can shape microbial communities in a variety of environments rang-

ing from the cystic fibrosis lung to the rhizosphere of dryland crops. For example, in the rhizosphere, phenazines can protect

plants from infection by pathogenic fungi. The redox activity of phenazines underpins their antibiotic activity, as well as provid-

ing pseudomonads with important physiological benefits. Our discovery that soil mycobacteria can catabolize phenazines and

thereby protect other organisms against phenazine toxicity suggests that phenazine degradation may influence turnover

in situ

The identification of genes involved in the degradation of phenazines opens the door to monitoring turnover in diverse environ-

ments, an essential process to consider when one is attempting to understand or control communities influenced by phenazines.

Received

7 September 2015

Accepted

5 October 2015

Published

27 October 2015

Citation

Costa KC, Bergkessel M, Saunders S, Korlach J, Newman DK. 2015. Enzymatic degradation of phenazines can generate energy and protect sensitive organisms from

toxicity. mBio 6(6):e01520-15. doi:10.1128/mBio.01520-15.

Editor

Frederick M. Ausubel, Massachusetts General Hospital

Creative Commons Attribution-Noncommercial-ShareAlike 3.0 Unported

license

, which permits unrestricted noncommercial use, distribution, and reproduction in any medium, provided the original author and source are credited.

Address

correspondence to Dianne K. Newman, dkn@caltech.edu.

P

seudomonas

spp. are i

mportant biocontrol agents that pro-

duce a variety of secreted metabolites that suppress disease

in the rhizosphere (1–4). Of these metabolites, phenazines are

an important subclass. Phenazines have long been studied due

to their antibiotic activities against diverse cell types as well as

their beneficial physiological roles for their producers (5). In

agricultural settings, the production of phenazines—particu-

larly phenazine-1-carboxylic acid (PCA) and phenazine-1-

carboxamide (PCN)—is thought to protect plants from colo-

nization and infection by pathogenic fungi (1–3). Phenazines

accumulate in the rhizosphere of dryland cereals, where they

have a half-life of 3.4 days (2). In addition to the rhizosphere,

phenazines are present and active in other environmental and

clinical contexts, such as crude oil and the lungs of patients

with the genetic disorder cystic fibrosis (CF) (6–8); however,

their turnover has not been measured in any of these systems.

While the relatively short half-life of phenazines in the rhizo-

sphere suggests that there are active mechanisms of removal, it

is unclear what they are.

Many natural phenazine compounds with a common nitrogen-

heterocyclic core have been described (Fig. 1A).

Pseudomonas

spp.

and other bacteria produce several phenazine derivatives with

diverse properties (9). The most abundant phenazines in

laboratory-grown

Pseudomonas aeruginosa

culture are PCA, PCN,

and pyocyanin (PYO). PCA is the precursor from which all other

phenazines are derived, PYO is produced by the action of two

enzymes (PhzM and PhzS) that modify PCA, and PCN is gener-

ated from PCA by the action of PhzH (10).

P. aeruginosa

can also

produce 1-hydroxyphenazine through the action of PhzS.

Phenazines benefit producing organisms in a variety of ways. In

P. aeruginosa

, phenazines are involved in anaerobic survival, iron

acquisition, signaling, and biofilm development (11–14). How-

ever, the redox properties of phenazines are harmful to other bac-

RESEARCH ARTICLE

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teria and eukaryotic organisms that are often found in association

with

Pseudomonas

spp. (15–17).

Phenazine toxicity differs depending on the phenazine type

and can change under various environmental conditions. For ex-

ample,

Caenorhabditis elegans

is more sensitive to PCA than PYO

at acidic pH, but the opposite is true at alkaline pH (15).

Phenazines can cause toxicity by producing reactive oxygen spe-

cies (ROS) and interfering with the respiratory electron transport

chain (16, 17). While defense against the toxic effects of

phenazines is generally thought to involve the induction of ROS

defense systems, the capacity to degrade or transform phenazines,

including PCA and PCN, has also been demonstrated (18–20). A

recent study showed changes to phenazines in mixed communi-

ties, where diffusion of phenazines between colonies of

P. aerugi-

nosa

and

Aspergillus fumigatus

results in several metabolic trans-

formations (but not removal) of the phenazines (21). Yet, in

analogy to what has been shown for acyl-homoserine lactone

quorum-sensing signal degradation (22, 23), it is important to

consider turnover processes in addition to chemical modifications

when one is seeking to understand the fate of phenazines in the

environment.

While the capacity to alter or degrade phenazines has been

demonstrated by microbes associated with

Pseudomonas

spp. in

natural communities (18–21), the genes responsible for this activ-

ity have been unknown. Here, we isolated phenazine-degrading

organisms and identified genes involved in the degradation of

three

Pseudomonas

-derived phenazines. We used these findings to

explore the effects of phenazine degradation on phenazine pro-

ducers (pseudomonads) and degraders (mycobacteria) and to de-

termine whether phenazine degradation can play a protective role

for other, phenazine-sensitive organisms. Our findings suggest

that the interactions between phenazines and phenazine degrad-

ers have the potential to tune the concentrations of different

phenazine types, and if phenazine degradation is active

in situ

,it

would be expected to impact microbial community structure.

RESULTS

Isolation of PCA-degrading organisms.

PCA is the precursor of

all phenazines produced by

Pseudomonas

spp. (9, 10). Therefore,

we reasoned that the ability to degrade PCA would be common

among organisms capable of degrading

Pseudomonas

-derived

phenazines. Soil was collected from 16 locations around the Cal-

ifornia Institute of Technology campus and the nearby San Ga-

briel mountains. Samples from six sites yielded isolates capable of

growing in medium with 2.5 mM PCA as the sole source of carbon

and energy (Fig. 1B). Among the isolates that were verified to

degrade PCA, all had similar colony morphology; thus, one isolate

from each site was randomly selected for follow-up studies. Partial

16S rRNA gene sequences were determined for each isolate and

phylogenetic trees were constructed. PCA-degrading organisms

grouped with two members of the genus

Mycobacterium

Myco-

bacterium fortuitum

(GenBank accession number NR_118883) and

Mycobacterium septicum

(accession number NR_042916), with

99% sequence identity (Fig. 1C). Strains CT6 (

M. fortuitum

like) and DKN1213 (

M. septicum

-like) were chosen as representa-

tive strains for further study.

Strains CT6 and DKN1213 can additionally degrade PYO.

While PCA is produced by all pseudomonads capable of

phenazine production, PYO is produced only by

P. aeruginosa

and

is the best-studied phenazine due to its clinical relevance. We

therefore sought to determine if our isolates were capable of de-

grading PYO. Because PYO is poorly soluble in aqueous solutions

at circumneutral pH, we monitored PYO degradation using mi-

cromolar— but physiologically relevant (24)— concentrations

with cultures grown in 10% LB-Tw (lysogeny broth with 0.05%

Tween 80) medium. Both CT6 and DKN1213 were capable of

FIG 1

Isolation, identification, and degradation phenotype of phenazine-degrading bacteria. (A) Structure of phenazines. For PCA, Y

COOH; for PCN, Y

CONH

; for PYO, X

and Y

OH. (B) Growth of representative isolates CT6 and DKN1213 with PCA as a sole source of carbon and energy.

P. aeruginosa

and

M. smegmatis

were both incapable of growth under this condition. (C) 16S rRNA gene dendrogram of various phenazine-degrading bacteria (red) and close

relatives. Organisms in blue were tested for phenazine degradation but showed no activity. (D) Growth and degradation of PYO by strains CT6 (red) and

DKN1213 (blue) supplied with 10% LB-Tw as a carbon source. Data are averages and standard deviations (SD) for three cultures. Solid lines, PYO concentration;

dashed lines, OD.

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degrading PCA under these conditions. Both strains were addi-

tionally capable of degrading PYO from a starting concentration

of 200

M to a final concentration of less than 20

M (the detec-

tion limit of our assay) (Fig. 1D) with a concomitant change of the

medium from blue to colorless. No change in medium color or

measured PYO was observed for the close relative

Mycobacterium

smegmatis

(see Fig. S1 in the supplemental material).

Genome sequence of strain CT6 and identification of candi-

date dioxygenases involved in PCA degradation.

Phenazines are

polycyclic, aromatic, nitrogen-containing heterocycles (Fig. 1A);

therefore, we hypothesized that the genes responsible for their

degradation likely encode dioxygenases, a family of enzymes

known to break down aromatics. Several observations lend cre-

dence to this notion. First, because the only source of carbon in

PCA that is sufficiently reduced to support growth is bound within

the ring, it must be cleaved; the carboxylate moiety alone cannot

support growth (Fig. 1A). Second, previous studies of the PCA

degradation pathway in

Sphingomonas wittichii

strain DP58, the

only other known PCA degrader, demonstrated that ring cleavage

is essential for PCA catabolism (25). Furthermore, in our cultures,

we never observed the accumulation of the unsubstituted core

phenazine molecule, despite the fact that our mycobacteria can

degrade this molecule (see Fig. S2 in the supplemental material);

its absence strongly suggests that the core phenazine ring is

cleaved. Finally, the fact that our mycobacteria can also use PCA as

a sole nitrogen source (see Fig. S3 in the supplemental material)

further indicates that ring cleavage must occur in order for nitro-

gen to be liberated from PCA (Fig. 1A). Accordingly, we focused

on identifying putative dioxygenase enzymes that might catalyze

PCA degradation.

The genome of

S. wittichii

strain DP58 has been sequenced and

contains 91 predicted dioxygenase genes, though none have been

identified as important for PCA degradation (19, 26). We sought

to identify candidate dioxygenase genes for PCA degradation by

comparative genomic analysis of

S. wittichii

DP58 and one of the

phenazine-degrading mycobacteria described here. The genome

of strain CT6 was sequenced to completion using the PacBio

single-molecule, real-time (SMRT) genome sequencing technol-

ogy. The strain CT6 genome is 6.25 Mbp and contains no plas-

mids. There are a predicted 6,051 protein-encoding genes accord-

ing to the RAST (Rapid Annotation using Subsystem Technology)

server, and the genome contains 35 to 47 dioxygenase genes as

annotated by RAST or the Bacterial Annotation System (BASys)

pipelines, respectively (27–29). Reciprocal best BLAST analysis of

annotated dioxygenase genes was performed between

S. wittichii

strain DP58 and strain CT6 (PCA degraders) or

M. smegmatis

-155 and strain CT6 to identify genes present in both PCA-

degrading organisms but absent from

M. smegmatis

. This

identified three predicted dioxygenase genes—XA26_16830,

XA26_16860, and XA26_16890 —that are co-oriented in an ~10-

kbp chromosomal locus in strain CT6 (Table 1; also, see Ta-

ble S1 in the supplemental material). Because the same set of genes

is present in both sequenced PCA-degrading organisms, we hy-

pothesized that these genes would be regulated specifically by

phenazines. Quantitatve reverse transcription-PCR (qRT-PCR)

analysis of the expression of putative dioxygenase genes in strain

CT6 demonstrated that these three genes, as well as a nearby

fourth gene predicted to encode a dioxygenase (XA26_16730), are

highly induced in the presence of phenazines (Fig. 2A). mRNA

abundance for these genes was increased

1,000-fold in the pres-

ence of both PCA and PYO after a 3-h exposure but not in the

presence of 9,10-anthraquinone-2,6-disulfonate (AQDS) or

methylene blue (MB), two other 3-ringed, aromatic, redox-active

molecules (13). AQDS and MB have midpoint redox potentials

that are lower and higher, respectively, than those of phenazines

(13) suggesting that a redox switch is not responsible for increased

expression. The mRNA abundance of two predicted dioxygenase

genes located distantly on the chromosome was unchanged in the

presence of any compounds tested.

Rhodococcus

strain JVH1 and

M. fortuitum

ATCC 6841 are

both capable of degrading phenazines.

A comparison of the ge-

nome of CT6 to that of the type strain

M. fortuitum

ATCC 6841

revealed the presence of the predicted PCA-degrading genes, and

this organism was verified to degrade both PCA and PYO (see

Fig. S1 in the supplemental material). The putative dioxygenases

predicted to be involved in PCA degradation were analyzed in the

Integrated Microbial Genomes/Expert Review (IMG/ER) web

server (30, 31), and

Rhodococcus

sp. strain JVH1 was also found to

contain these genes (32, 33). Based on their presence, we predicted

that

Rhodococcus

sp. strain JVH1 would be capable of PCA degra-

dation. When suspended at high density (optical density at 600

nm [OD

600

] of ~2 to 3), JVH1 degraded 200

M PCA to ~50% of

the starting concentration in 48 h—a rate of degradation signifi-

cantly lower than that observed for mycobacteria. No activity was

TABLE 1

Genes important to this study

CT6

M. fortuitum

ATCC 6841

Rhodococcus

JVH1

Sphingomonas

DP58

XA26 gene no.

RAST annotation

MFORT gene no.

% ID

Gene GI no.

% ID

MSEDRAFT gene no.

% ID

16600

3-Hydroxy-4-oxoquinaldine 2,4-dioxygenase

16204

100

00241

16610

Salicylate hydroxylase

16209

497117731

05232

16620

Hypothetical protein

16214

760099504

02403

16640

4-Hydroxyphenylacetate 3-monooxygenase

16224

100

396930211

16730

Ortho-halobenzoate 1,2-dioxygenase

16269

497121742

05237

16830

2,3-Dihydroxybiphenyl 1,2-dioxygenase

16319

497121763

04471

16860

Large subunit naph/bph dioxygenase

16334

100

497121769

04467

16890

2,3-Dihydroxybiphenyl 1,2-dioxygenase

16349

497121775

04486

16960

Amidase

30529

497116141

05146

16980

Probable oxidoreductase

14347

497109126

04185

16990

Hypothetical protein

14352

An expanded version of this table is presented as Table S1 in the supplemental material. ID, identity.

naph/bph, naphthalene/biphenyl.

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seen in

Rhodococcus

sp. strain RHA1, the closest sequenced rela-

tive that lacks the genes of interest (Fig. 2B). No degradation ac-

tivity was observed with PYO for either strain.

Allelic replacement of candidate dioxygenase homologs

from

M. fortuitum

ATCC6841abolishestheabilitytogrowwith

PCA as the sole carbon source.

The presence of the same putative

dioxygenase genes in all PCA degraders and their phenazine-

specific regulation in strain CT6 strongly suggest an involvement

in PCA degradation. We used mutagenesis to test this hypothesis.

Attempts to genetically manipulate strain CT6 were ineffective,

but a similar approach succeeded in the

M. fortuitum

type strain. A

recombineering approach was employed to create mutations

in the four putative dioxygenase genes

(MFORT_16269, MFORT_

16319, MFORT_16334, and MFORT_16349) (Table 1) (34), and

each gene was replaced with a gentamicin resistance cassette.

Gentamicin-resistant colonies with disruptions in each gene were

streaked onto agar medium with 2.5 mM PCA as the sole carbon

source. None of the mutants grew under these conditions, indi-

cating that the mutated genes are essential for growth with PCA as

a carbon source (Fig. 2C). High-performance liquid chromatog-

raphy (HPLC) analysis of supernatants from cultures grown on

LB medium with 100

M PCA revealed the loss of PCA from

16319::Gm

16334::Gm

, and

16349::Gm

strains but not

from the

16269::Gm

strain (see Fig. S4 in the supplemental ma

terial). This suggests that the product of MFORT_16269 is neces-

sary to catalyze PCA degradation, whereas the other genes are

involved in a downstream reaction (these mutants cannot use

PCA as a carbon source [Fig. 2C] but still remove it from culture

supernatants [see Fig. S4 in the supplemental material]). None of

these mutants had a defect in PYO removal.

Notably, only the putative single-subunit, ring-cleavage dioxy-

genase genes, MFORT_16319 and MFORT_16349, could be com-

plemented in

trans

(see Fig. S5 in the supplemental material). It is

unclear why the putative multisubunit, ring-hydroxylating dioxy-

genase genes MFORT_16269 and MFORT_16334 could not be

complemented, but perhaps these enzymes are nonfunctional

when overexpressed. The position of the gentamicin resistance

cassette on the chromosome was verified by sequencing for each

mutation; however, we cannot rule out the possibility that a

secondary site mutation resulted in the phenotype for the

MFORT_16269 and MFORT_16334 mutants.

RNA-Seq and mutagenesis identifies a small hypothetical

protein that is necessary for PYO degradation.

Because the four

predicted dioxygenase genes were expressed specifically in the

presence of phenazines, we took a transcriptome sequencing

(RNA-Seq) approach to identify candidate genes important to

PYO degradation. Strain CT6 was used for RNA-Seq because the

genome of this organism has been closed, whereas the genome of

ATCC 6841 exists as 82 contigs (35). As

expected, the genes im-

portant for PCA-dependent growth showed increased mRNA

abundance after a 20-min exposure to either PCA or PYO

(Fig. 3A). In fact, the entire region of the CT6 genome that

shares homology with

Rhodococcus

sp. strain JVH1 (an organ-

ism that can degrade only PCA) was highly expressed in the

presence of either PCA or PYO. Additionally, genes flanking

this region had increased mRNA abundance and were induced

to a greater extent by PYO.

A mutant lacking the entire ~40-kb region (

40kb::Gm

)of

the genome induced by phenazines was constructed in

M. for-

tuitum

and found to completely lack the ability to degrade

phenazines, including PYO (Fig. 3B). This locus contains genes

for an additional predicted dioxygenase (MFORT_16204

[XA26_16600]) and a monooxygenase (MFORT_16224 [XA26_

16640]) in one of the flanking regions; however, mutants with

mutations in both genes were still capable of PYO degradation

(Fig. 3B). Of note is that

16224::Gm

degrades PYO more

slowly than the wild type (WT), suggesting a possible role in a

downstream reaction in the PYO degradation pathway. A mu-

tant missing a three-gene operon in the other flanking region

(MFORT_14352 to MFORT_30529 [XA26_16990 to XA26_

16960]) lacked the ability to degrade PYO. A single gene mu-

tant lacking MFORT_14352, annotated as a hypothetical pro-

tein, was deficient in PYO degradation; additionally,

expression of this gene in

Rhodococcus

sp. strain JVH1 allowed

PYO degradation by this strain (see Fig. S6 in the supplemental

material). Therefore, MFORT_14352 is necessary for the first

step of PYO degradation and may be sufficient for the removal

of this phenazine from the medium.

FIG 2

Evidence for the involvement of four putative dioxygenase genes in the

breakdown of PCA. (A) qRT-PCR of genes from cultures exposed to various

redox-active small molecules. Data are normalized internally to

rpoA

and to a

control with no added small molecule. Data represent at least duplicates, and

error bars show one SD around the mean for triplicate cultures. (B) Phenazine

degradation by

Rhodococcus

strains JVH1 and RHA1. Data are averages and SD

for three independent cultures. (C) Growth of individual gene mutants with

PCA as a sole carbon source.

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M. fortuitum

alters the phenazine pool in coculture with

Pseudomonas

spp.

M. fortuitum

is ubiquitous and found in envi-

ronments where phenazine-producing pseudomonads are also

present (36, 37). Using a subset of the mutants described above,

we sought to determine whether a mixed culture of

M. fortuitum

with a pseudomonad would lead to alterations in the phenazine

pool.

M. fortuitum

and several mutants were inoculated into LB

medium with

P. aeruginosa

PA14, and culture supernatants were

analyzed by HPLC after 24 h. The

40kb::Gm

mutant was used as

a control to determine the amount of phenazines produced by

PA14 under coculture conditions. The WT

M. fortuitum

strain

decreased the abundance of both PYO (~50% decreased) and

PCN (~90% decreased) but not PCA in coculture with PA14

(Fig. 4A). This level of phenazine degradation impacted neither

the rate of

P. aeruginosa

growth nor its transcription of the

phenazine biosynthesis genes. As expected, the PYO degradation

mutant,

14352::Gm

, decreased the abundance of PCN but not

PYO. The mutant

30529::Gm

decreased PYO but not PCN, sug

gesting that MFORT_30529 —annotated as an amidase—may

convert PCN to PCA. In fact, cocultures that are proficient for

PCN degradation seem to have higher PCA concentrations in

their supernatants, supporting this hypothesis (Fig. 4A; also, see

Fig. S6 in the supplemental material). The PCN degradation phe-

notype of

30529::Gm

was assayed in monoculture, and this

strain was confirmed to have a specific defect in the removal of

PCN from culture supernatants (see Fig. S7 in the supplemental

material); additionally, expression of MFORT_30529 in

Rhodo-

coccus

sp. strain JVH1 conferred the ability to degrade PCN to this

organism (see Fig. S6).

Because

M. fortuitum

in coculture with PA14 prioritizes PYO

and PCN removal from the medium, to determine whether PCA

could also be degraded in coculture, we mixed

M. fortuitum

and

FIG 3

Gene expression and mutational analysis of PYO degradation. (A) mRNA abundance of strain CT6 genes after 20-min exposure to PCA or PYO. Values

are given as log

fold change versus a mock treatment. An ~40-kb region of the genome shows a large change in gene expression in response to phenazines. Strain

CT6 gene numbers are listed on the

axis, with genes on the plus strand listed on the top row and genes on the minus strand listed on the bottom row. Genes

highlighted in red are present in the genome of

Rhodococcus

strain JVH1 (Table 1; also, see Table S1 in the supplemental material). Blue points represent relative

mRNA abundance after PYO exposure, and orange points correspond to PCA exposure. (B) Growth and PYO degradation characteristics of

M. fortuitum

mutants grown in 10% LB-Tw medium supplemented with 200

M PYO. All mutants could grow with 10% LB as the carbon source, but only a subset displayed

a defect in PYO degradation. Data are averages and SD for three cultures. RNA-Seq data for the entire chromosome can be found in Dataset S1 in the

supplemental material.

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Pseudomonas fluorescens

2-79, an organism that produces PCA as

its only phenazine. WT

M. fortuitum

was capable of removing

~50% of the PCA from culture supernatants compared to the

40kb::Gm

control (Fig. 4B). As expected,

16269::Gm

was de

fective for PCA removal. For unknown reasons,

30529::Gm

was

also defective for PCA removal in the coculture condition.

PYO degradation is protective to other organisms in

coculture with

M. fortuitum

Alterations in the abundance of

phenazines in cocultures of

M. fortuitum

and

Pseudomonas

spp.

demonstrate that phenazine degradation can influence the con-

centration of structurally diverse phenazines in the coculture en-

vironment. Because phenazines can function as antimicrobials

(16, 17), it is ecologically important to consider whether degrada-

tion might impact other members of a microbial community. We

hypothesized that PYO degradation would be protective to organ-

isms that are otherwise susceptible. PYO survival was assayed in

cocultures of

M. fortuitum

mixed with diverse bacteria that have a

range of sensitivity to different phenazines:

Staphylococcus aureus

Agrobacterium tumefaciens

Shewanella oneidensis

, and

Escherichia

coli

(16, 17, 38, 39). After 24 h of incubation in monoculture in the

presence of 100

M PYO,

S. aureus

E. coli

, and

S. oneidensis

were

all inhibited by PYO to various degrees (Fig. 5). In all three cases,

coculture with WT

M. fortuitum

rescued growth of these organ-

isms, though not always to the same extent as growth in monocul-

ture in the absence of PYO. A PYO degradation mutant provided

no protection.

A. tumefaciens

was resistant to PYO, consistent

with previous reports (38), and coculture had little effect on this

organism.

DISCUSSION

Phenazines can shape both microbial community composition

and the chemistry of the environment; however, while much is

known about their biosynthesis, regulation and physiological

functions (11–14), little is known about their degradation. Previ-

ously,

S. wittichii

DP58 was shown to be capable of degrading

PCA, yet the genes catalyzing this process were not identified (25).

Here, we identified genes involved in the degradation of multiple

phenazines in members of the

Mycobacterium fortuitum

complex.

The identification of conserved degradation genes in mycobacte-

ria not only broadens the phylogenetic diversity of this activity to

include members of both the

Proteobacteria

and

Actinobacteria

phyla it also suggests that the enzymes catalyzing this activity may

be widespread.

Mycobacteria are ubiquitous, and

Mycobacterium

spp. and

Pseudomonas

spp. are commonly reported to be present in the

same types of environments, including soil, crude oil, and the

lungs of patients with CF (6–8). It may thus not be surprising that

one organism has evolved the capacity to utilize an excreted prod-

uct of the other. A common gene cluster that appears to be essen-

tial for PCA degradation is shared between the genomes of

M. for-

tuitum

ATCC 6841,

Rhodococcus

sp. strain JVH1, and strain CT6.

Notably, the same four putative dioxygenases display increased

mRNA abundance in strain CT6 in the presence of both PCA and

PYO.

Rhodococcus

strain JVH1 is incapable of PYO degradation

yet possesses close homologs of each of these genes. This suggests

that PCA and PYO may share a degradation intermediate. In

P. aeruginosa

, PYO is produced from PCA via the action of PhzS

and PhzM (10). One possibility is that strains CT6 and DKN1213

first degrade PYO to PCA and that this product leads to the in-

creased mRNA abundance for PCA-specific genes. A second pos-

sibility is that PCA and PYO may be converted to an intermediate

FIG 4

Phenazine degradation by

M. fortuitum

strains grown in coculture

with

Pseudomonas

spp. Cocultures were grown for 24 h, and phenazine con-

centrations were measured by HPLC.

40 kb::Gm

is defective in phenazine

degradation, so PYO, PCN, and PCA levels in a coculture with this strain were

arbitrarily set to 100%. (A) Phenazine levels in a coculture between the indi-

cated mutant and

P. aeruginosa

PA14. (B) PCA levels in a coculture between

the indicated mutant and

P. fluorescens

2-79. Data are averages and SD from

three experiments.

FIG 5

PYO degradation protects sensitive organisms.

E. coli

S. oneidensis

S. aureus

, and

A. tumefaciens

were plated after coculture with WT

M. fortuitum

14352::Gm

(

) in the presence of 100

M PYO. Survival of these organisms in monoculture with 0 or 100

M PYO is included to verify their sensitivity to PYO.

Sensitive organisms had increased cell density after incubation with the WT, suggesting a protective effect for PYO degradation.

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that is further degraded through the action of MFORT_16269,

MFORT_16319, MFORT_16334, and MFORT_16349 or a subset

of these. A small (162-amino-acid) protein is sufficient to catalyze

PYO degradation, and a predicted amidase is likely required for

the first step in PCN breakdown. An amidase activity suggests that

PCN is first converted to PCA before further breakdown takes

place. Elucidation of the complete pathway(s) of PCA, PCN, and

PYO degradation awaits future research, yet this study provides an

important step in that direction.

What consequences might the degradation of different

phenazines have on

Pseudomonas

spp.? Though we did not ob-

serve significant phenotypic consequences of phenazine degrada-

tion on

P. aeruginosa

under standard laboratory growth condi-

tions, it is possible that phenazine degradation may decrease the

fitness of phenazine-producing organisms in natural or engi-

neered environments. Each

Pseudomonas

-derived phenazine has

unique redox and chemical properties (13, 40) and a distinct im-

pact on the physiology of its producer (9, 41). For example, PYO is

important for biofilm maturation in

P. aeruginosa

(42), and PCA

rapidly reduces ferric iron, facilitating iron acquisition (12).

Phenazine toxicity to other organisms can benefit

Pseudomonas

spp. by inhibiting competitors. Finally, phenazines are terminal

signaling factors in the quorum-sensing network of pseudomon-

ads (14). In many cases, the impact of molecular signals is

concentration dependent, but how degradation influences the ac-

cumulation and fate of bacterial signals is generally poorly char-

acterized. However, as shown by work on acyl-homoserine lac-

tone degradation, the ability to control signal accumulation can

significantly affect the behavior of microbes responsive to that

signal if it is consumed below a certain threshold (22, 23). We thus

speculate that controlling the distribution of phenazines via deg-

radation might similarly attenuate the ability of

Pseudomonas

spe-

cies to dominate environments where phenazine cycling is bene-

ficial (e.g., hypoxic or anoxic habitats) or provides a competitive

advantage.

Just as the degradation of phenazine metabolites would be pre-

dicted to impact the fitness of phenazine producers, we might also

anticipate a more general effect on ecosystem diversity.

P. aerugi-

nosa

interactions with plants highlight the potential implications

of this interaction on the outcome of two different fungal infec-

tions. PYO enhances susceptibility of rice to

Rhizoctonia solani

but

leads to increased resistance to

Magnaporthe grisea

(43); the pres-

ence of a PYO-degrading organism could significantly alter the

outcome of infection in each of these cases. While there is evidence

of phenazine turnover and degradation in the rhizosphere (2, 19),

phenazines also play important roles in other environments where

these processes have not been measured or observed. The presence

of phenazines is negatively correlated with microbial species rich-

ness in both laboratory enrichment cultures and the lungs of pa-

tients with CF (6–8, 44). What role might natural or stimulated

degradation play in modulating phenazine levels and microbial

community development in these and other contexts? Given that

phenazine degradation can impact the fitness of organisms by

promoting growth and minimizing toxicity, it is temting to spec-

ulate that these activities may be important in natural environ-

ments. Future work will determine whether this is, in fact, the case.

MATERIALS AND METHODS

Strains, media, and growth conditions.

Strains used in this study are

listed in Table S2 in the supplemental material. PCA-degrading organisms

were isolated by inoculating 5 ml of PCA medium with 100 mg of soil

collected on and around the California Institute of Technology campus

and growing at 30°C with shaking. PCA medium was composed of mini-

mal medium containing (per liter) KH

(1.93 g), K

HPO

(6.24 g),

NaCl (2.5 g), MgSO

(0.12 g), NH

Cl (0.5 g) and supplemented with PCA

(0.56 g). Trace minerals were also included (45). Cultures were allowed to

grow for 5 to 8 days before transfer to fresh PCA medium. After 3 rounds

of growth in PCA liquid medium, cultures were streaked on plates con-

taining PCA medium amended with 1.5% Bacto agar. After isolation,

strains were routinely grown in lysogeny broth with 0.05% Tween 80

(LB-Tw). PCA and PCN were purchased from Princeton Biomolecular

Research Inc., and PYO was purified from cultures of

P. aeruginosa

strain

PA14 grown with 40 mM succinate as the sole carbon source using organic

extraction into dichloromethane as described previously (14).

Growth curves were performed in 10% LB-Tw medium supplemented

with 200

M PCA, PYO, or PCN. Cultures were grown to mid-

exponential phase and diluted to an OD of ~0.1 to 0.2 for growth analysis.

Growth curves were carried out in 96-well plate format on a BioTek Syn-

ergy 4 microplate reader (BioTek, Inc.) set to 30°C and medium agitation.

Ninety-six-well plates were sealed with sterile adhesive film to prevent

medium evaporation. OD was monitored at 500 nm; PCA and PCN were

monitored at 370 nm, and PYO was monitored at 310 nm. Standards were

included to calculate the concentrations of each phenazine. PCA and PYO

degradation by

Rhodococcus

was performed at an OD

600

of ~2 to 3.

Rho

dococcus

spp. were suspended in 10% LB medium with 200

M PCA or

PYO in a volume of 5 ml. A 500-

l portion of culture supernatant was

collected by centrifugation at various time points and analyzed on a plate

reader.

Coculture experiments were carried out in LB with various

M. fortui-

tum

strains at a starting OD of 0.15. Pseudomonads were inoculated to an

OD of 0.05 and other organisms to 0.001. Cocultures were grown for 24 h

in all cases except for

A. tumefaciens

(48 h). To test the protective effects of

PYO degradation, 100

M PYO was included, and cultures were plated on

LB at the end of growth to enumerate CFU (all organisms formed colonies

in 1 to 2 days versus 3 to 5 days for

M. fortuitum

, so selective medium was

not necessary). To analyze coculture supernatants, samples were diluted

1:10, sterilized by filtration on a 0.2-

m filter and analyzed by HPLC as

described previously (12) using the same method but a flow rate of 950

min

Phylogenetic analysis.

16S rRNA genes were amplified using primers

9bf and 1512uR and sequenced with 9bf and 519uF (46, 47).

Relatives

were found using BLAST (

http://blast.ncbi.nlm.nih.gov

) and collected

for

analysis. All phylogenetic analyses were performed using the phylog-

eny.fr web server and further refined using Interactive Tree Of Life (iTOL)

(48–51).

DNA extraction, genome sequencing, gene annotation and analysis.

DNA extraction of strain CT6 was performed using a modified protocol

from (52). Fifty milliliters of mid-exponential-phase culture was har-

vested and stored at

80°C before DNA extraction. Pellets were thawed at

80°C for 30 min and suspended in an equal volume of chloroform-

methanol (2:1) for 1 h with periodic shaking at room temperature. This

mix was centrifuged for 20 min at 2,500

, and the organic and aqueous

phases were discarded without disturbing the cell material. The cell pellet

was dried for 10 min at 55°C, suspended in 550

l of 100 mM Tris

(pH 9.5), and incubated overnight at 37°C with lysozyme (final concen-

tration, 0.5 mg ml

). After lysozyme treatment, SDS (final concentra

tion, 1%) and proteinase K (final concentration, 0.75 mg ml

) were

added, and the tube was left at 60°C for 1 h with periodic mixing. During

the last 15 min of incubation, 100

l each of 5 M NaCl and cetyltrimeth-

ylammonium bromide (CTAB) solution (10% CTAB solution in 0.7 M

NaCl) was added. DNA was

extracted with 1 vol of chloroform-isoamyl

alcohol (24:1), and the aqueous phase was treated with RNase A. RNA-

free DNA was ethanol precipitated and stored at

80°C before se-

quencing.

SMRT sequencing was performed using the Pacific Biosciences RS II

Degradation of Phenazines by Mycobacteria

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platform with 10-kb libraries and P5/C3 chemistry using the standard

protocols. The

de novo

genome assembly, using HGAP v2 (53), resulted in

a single circular contig. The

genome of strain CT6 was annotated using

RAST and BASys (27–29). Genomes were analyzed using BLAST

(

http://blast.ncbi.nlm.nih.gov

) and the Integrated Microbial Genomes/

Expert

Review (IMG/ER) web server (

https://img.jgi.doe.gov/er/

) (26, 30,

31,

35).

RNA extraction, qRT-PCR, and RNA-Seq.

To generate material for

qRT-PCR analysis, overnight cultures were diluted 1/50 in LB-Tw and

grown overnight to an OD

600

of ~0.3. At this point, PCA, PYO, AQDS, or

MB was added to a final concentration of 200

M. An equivalent volume

of water was added to a set of cultures as a control. After 3 h, cell material

was collected, flash frozen, and stored at

80°C until RNA extraction.

Frozen cell pellets were suspended in 350

l AES buffer (50 mM sodium

acetate [pH 5.3], 10 mM EDTA, 1% SDS) and 350

l acid phenol-

chloroform (5:1; pH 4.5), and 200

l glass beads (

106

m) were added.

Samples were homogenized in an analog Disruptor Genie (Scientific In-

dustries, Inc.) four times for 30-s each with a 30-s interval on ice between

disruptions. Glass beads were removed by centrifugation, and liquid was

transferred to a heavy phase lock tube containing 300

l chloroform. The

phase lock tube was centrifuged for 5 min at 12,000

. The aqueous

phase was extracted with an additional 300

l chloroform and transferred

to a microcentrifuge tube containing 40

l 3M sodium acetate (pH 5.2),

and RNA was alcohol precipitated and suspended in 100

O. This

crude RNA extract was cleaned using an RNeasy kit (Qiagen, Inc.) with a

modified protocol with optional on-column DNase treatment, as de-

scribed previously (54). RNA was additionally treated with Turbo DNA-

free using the manufacturer’s directions (Life Technologies, Inc.).

Purified RNA was converted to cDNA using an iScript cDNA synthesis

kit and following the manufacturer’s directions (Bio-Rad, Inc.). cDNA

was used as a template for qPCR using iTaq universal SYBR green Super-

mix (Bio-Rad, Inc.) on a 7500 fast real-time PCR system (Applied Biosys-

tems, Inc.). Samples were analyzed in at least duplicate, and the signal

from each treatment (PCA, PYO, AQDS, or MB) was first normalized to a

water-only control using the following equation: relative expression

[

(sample)

(control)]

where

is the calculated efficiency for a given

primer pair. Data were then standardized to

rpoA

. qRT-PCR primers are

listed in Table S2 in the supplemental material.

For RNA-Seq, RNA was extracted after a 20-min exposure to 200

PCA, PYO, or water. rRNA was depleted with the magnetic RiboZero kit

for Gram-negative bacteria (Epicentre). A library was prepared using the

NEBNext mRNA library prep master mix set for Illumina (NEB). Se-

quencing was performed at the Millard and Muriel Jacobs Genetics and

Genomics Laboratory at the California Institute of Technology to a depth

of 12 to 15 million reads on an Illumina HiSeq2500 and processed using

the Illumina HiSeq control software (HCS version 2.0). Low-quality bases

were removed using Trimmomatic (LEADING:27 TRAILING:27 SLID-

INGWINDOW:4:20 MINLEN:35) (55), mapped using Bowtie (56), and

sorted with SAMtools (57). The number of reads per locus was calculated

with easyRNAseq (58). The .gff file was generated using the RAST anno-

tations (27, 28). Reads were normalized by number and compared to the

water control. RNA-Seq results can be found in Dataset S1 in the supple-

mental material.

Generation of mutants.

M. fortuitum

ATCC 6841 was used for mu-

tagenesis using a recombineering protocol (34, 59, 60). Plasmids and

primers are listed in Table S2 in the supplemental material.

M. fortuitum

ATCC 6841 was electroporated with pJV53 using a protocol established

for

M. smegmatis

to make a recombineering strain (34, 59). Cultures were

plated on LB plates with 100

gml

kanamycin and allowed to grow for

3 to 5 days.

M. fortuitum

ATCC 6841 pJV53 was grown overnight in

LB-Tw to an OD

600

of 0.5 to 1, induced for 3 h with 0.2% acetamide,

washed and suspended in 10% glycerol as described previously, and fro-

zen at

80°C (34).

Linear DNA constructs for generating allele replacement mutants

were generated by PCR amplifying genomic regions flanking the target

gene and the gentamicin resistance cassette from pMQ30 (61). PCR prod-

ucts were treated with XbaI and NotI (New England Biolabs) when nec-

essary (see Table S2 in the supplemental material) and T4 ligase (Invitro-

gen). Constructs were PCR amplified and electroporated into acetamide

induced

M. fortuitum

ATCC 6841 pJV53 and outgrown overnight in

LB-Tw medium at 30°C. Allele replacement mutants were selected on LB

plates with 100

gml

gentamicin at 30°C. The position of the gentami

cin cassette on the chromosome of single-gene-deletion mutants was con-

firmed by PCR and sequencing.

To complement mutations in

trans

, cultures were cured of pJV53 by

daily passage at 37°C for 5 days and streak purified. Colonies that had lost

pJV53 and by extension kanamycin resistance were identified by patch

plating on LB and on LB supplemented with kanamycin. Mutants cured of

pJV53 were subsequently transformed with plasmid pSD5 (60) bearing

the appropriate gene to complement the mutation.

Nucleotide sequence accession number.

The complete genome se-

quence of strain CT6 has been deposited in GenBank (accession number

CP011269).

SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at

http://mbio.asm.org/

lookup/suppl/doi:10.1128/mBio.01520-15/-/DCSupplemental

Dataset

S1, XLSX file, 0.4 MB.

Figure S1, TIF file, 2.8 MB.

Figure S2, TIF file, 2.8 MB.

Figure S3, TIF file, 2.8 MB.

Figure S4, TIF file, 2.7 MB.

Figure S5, TIF file, 2.7 MB.

Figure S6, TIF file, 2.7 MB.

Figure S7, TIF file, 2.8 MB.

Table S1, DOCX file, 0.1 MB.

Table S2, DOCX file, 0.1 MB.

ACKNOWLEDGMENTS

We thank Jon Van Hamme and Lindsay Eltis for providing

Rhodococcus

strains JVH1 and RHA1. We also thank members of the Newman lab for

helpful feedback.

K.C.C. was supported by a Ruth L. Kirschstein National Research Ser-

vice Award F32 from the NIH, award no. F32AI112248. This work was

further supported by the Howard Hughes Medical Institute (HHMI) and

the Millard and Muriel Genetics and Genomics Laboratory at the Califor-

nia Institute of Technology; D.K.N. is an HHMI investigator.

REFERENCES

Mavrodi DV, Mavrodi OV, Parejko JA, Bonsall RF, Kwak YS, Paulitz

TC, Thomashow LS, Weller DM

. 2012. Accumulation of the antibi-

otic phenazine-1-carboxylic acid in the rhizosphere of dryland cereals.

Appl Environ Microbiol

78:

804 – 812.

http://dx.doi.org/10.1128/

AEM.06784-11

Mavrodi

DV, Parejko JA, Mavrodi OV, Kwak Y, Weller DM, Blanken-

feldt W, Thomashow LS

. 2013. Recent insights into the diversity, fre-

quency and ecological roles of phenazines in fluorescent

Pseudomonas

spp. Environ Microbiol

15:

675– 686.

http://dx.doi.org/10.1111/j.1462

-2920.2012.02846.x

Puopolo

G, Masi M, Raio A, Andolfi A, Zoina A, Cimmino A, Evidente

. 2013. Insights on the susceptibility of plant pathogenic fungi to

phenazine-1-carboxylic acid and its chemical derivatives. Nat Prod Res

27:

956 –966.

http://dx.doi.org/10.1080/14786419.2012.696257

Keel

C, Schnider U, Maurhofer M, Voisard C, Laville J, Burger U,

Wirthner P, Haas D, Défago G

. 1992. Suppression of root diseases by

Pseudomonas fluorescens

CHA0: importance of the bacterial secondary

metabolite 2,4-diacetylphloroglucinol. Mol Plant Microbe Interact

4 –13.

http://dx.doi.org/10.1094/MPMI-5-004

Grahl

N, Kern SE, Newman DK, Hogan DA

. 2013. The yin and yang of

phenazine physiology, p 43– 69. In Chincholkar S, Thomashow L (ed),

Microbial phenazines. Springer, Berlin, Germany.

Norman RS, Moeller P, McDonald TJ, Morris PJ

. 2004. Effect of pyo-

cyanin on a crude-oil-degrading microbial community. Appl Environ Mi-

Costa et al.

®

mbio.asm.org

November/December 2015 Volume 6 Issue 6 e01520-15

mbio.asm.org

on January 13, 2016 - Published by

mbio.asm.org

Downloaded from

crobiol

70:

4004 – 4011.

http://dx.doi.org/10.1128/AEM.70.7.4004

-4011.2004

Bange

FC, Kirschner P, Bottger EC

. 1999. Recovery of mycobacteria

from patients with cystic fibrosis. J Clin Microbiol

37:

3761–3763.

Torrens JK, Dawkins P, Conway SP, Moya E

. 1998. Non-tuberculous

mycobacteria in cystic fibrosis. Thorax

53:

182–185.

http://dx.doi.org/

10.1136/thx.53.3.182

Mavrodi

DV, Blankenfeldt W, Thomashow LS

. 2006. Phenazine com-

pounds in fluorescent

Pseudomonas

spp. biosynthesis and regulation.

Annu Rev Phytopathol

44:

417– 445.

http://dx.doi.org/10.1146/

annurev.phyto.44.013106.145710

10.

Mavrodi

DV, Bonsall RF, Delaney SM, Soule MJ, Phillips G, Thom-

ashow LS

. 2001. Functional analysis of genes for biosynthesis of pyocyanin

and phenazine-1-carboxamide from

Pseudomonas aeruginosa

PAO1. J

Bacteriol

183:

6454 – 6465.

http://dx.doi.org/10.1128/JB.183.21.6454

-6465.2001

11.

Glasser

NR, Kern SE, Newman DK

. 2014. Phenazine redox cycling en-

hances anaerobic survival in

Pseudomonas aeruginosa

by facilitating gen-

eration of ATP and a proton motive force. Mol Microbiol

92:

399 – 412.

http://dx.doi.org/10.1111/mmi.12566

12.

Wang

Y, Wilks JC, Danhorn T, Ramos I, Croal L, Newman DK

. 2011.

Phenazine-1-carboxylic acid promotes bacterial biofilm development via

ferrous iron acquisition. J Bacteriol

193:

3606 –3617.

http://dx.doi.org/

10.1128/JB.00396-11

13.

Wang

Y, Kern SE, Newman DK

. 2010. Endogenous phenazine antibiotics

promote anaerobic survival of

Pseudomonas aeruginosa

via extracellular

electron transfer. J Bacteriol

192:

365–369.

http://dx.doi.org/10.1128/

JB.01188-09

14.

Dietrich

LEP, Price-Whelan A, Petersen A, Whiteley M, Newman DK

2006. The phenazine pyocyanin is a terminal signalling factor in the quo-

rum sensing network of

Pseudomonas aeruginosa

. Mol Microbiol

61:

1308 –1321.

http://dx.doi.org/10.1111/j.1365-2958.2006.05306.x

15.

Cezairliyan

B, Vinayavekhin N, Grenfell-Lee D, Yuen GJ, Saghatelian A,

Ausubel FM

. 2013. Identification of

Pseudomonas aeruginosa

phenazines

that kill

Caenorhabditis elegans

. PLoS Pathog

e1003101.

http://

dx.doi.org/10.1371/journal.ppat.1003101

16.

Voggu

L, Schlag S, Biswas R, Rosenstein R, Rausch C, Gotz F

. 2006.

Microevolution of cytochrome

oxidase in staphylococci and its impli-

cation in resistance to respiratory toxins released by

Pseudomonas

. J Bac-

teriol

188:

8079 – 8086.

http://dx.doi.org/10.1128/JB.00858-06

17.

Hassett

DJ, Charniga L, Bean K, Ohman DE, Cohen MS

. 1992. Response

Pseudomonas aeruginosa

to pyocyanin: mechanisms of resistance, anti-

oxidant defenses, and demonstration of a manganese-cofactored superox-

ide dismutase. Infect Immun

60:

328 –336.

18.

Hill JC, Johnson GT

. 1969. Microbial transformation of phenazines by

Aspergillus sclerotiorum

. Mycologia

61:

452– 467.

http://dx.doi.org/

10.2307/3757234

19.

Yang

Z, Wang W, Jin Y, Hu H, Zhang X, Xu Y

. 2007. Isolation,

identification, and degradation characteristics of phenazine-1-carboxylic

acid-degrading strain

Sphingomonas

sp. DP58. Curr Microbiol

55:

284 –287.

http://dx.doi.org/10.1007/s00284-006-0522-7

20.

Willumsen

PA, Johansen JE, Karlson U, Hansen BM

. 2005. Isolation and

taxonomic affiliation of N-heterocyclic aromatic hydrocarbon-

transforming bacteria. Appl Microbiol Biotechnol

67:

420 – 428.

http://

dx.doi.org/10.1007/s00253-004-1799-8

21.

Moree

WJ, Phelan VV, Wu CH, Bandeira N, Cornett DS, Duggan BM,

Dorrestein PC

. 2012. Interkingdom metabolic transformations captured

by microbial imaging mass spectrometry. Proc Natl Acad SciUSA

109:

13811–13816.

http://dx.doi.org/10.1073/pnas.1206855109

22.

Dong

Y, Wang L, Xu J, Zhang H, Zhang X, Zhang L

. 2001. Quenching

quorum-sensing-dependent bacterial infection by an N-acyl homoserine

lactonase. Nature

411:

813– 817.

http://dx.doi.org/10.1038/35081101

23.

Wang

YJ, Leadbetter JR

. 2005. Rapid acyl-homoserine lactone quorum

signal biodegradation in diverse soils. Appl Environ Microbiol

71:

1291–1299.

http://dx.doi.org/10.1128/AEM.71.3.1291-1299.2005

24.

Price-Whelan

A, Dietrich LEP, Newman DK

. 2007. Pyocyanin alters

redox homeostasis and carbon flux through central metabolic pathways in

Pseudomonas aeruginosa

PA14. J Bacteriol

189:

6372– 6381.

http://

dx.doi.org/10.1128/JB.00505-07

25.

Chen

K, Hu H, Wang W, Zhang X, Xu Y

. 2008. Metabolic degradation

of phenazine-1-carboxylic acid by the strain

Sphingomonas

sp. DP58: the

identification of two metabolites. Biodegradation

19:

659 – 667.

http://

dx.doi.org/10.1007/s10532-007-9171-1

26.

Z, Shen X, Hu H, Wang W, Peng H, Xu P, Zhang X

. 2012. Genome

sequence of

Sphingomonas wittichii

DP58, the first reported phenazine-1-

carboxylic acid-degrading strain. J Bacteriol

194:

3535–3536.

http://

dx.doi.org/10.1128/JB.00330-12

27.

Overbeek

R, Olson R, Pusch GD, Olsen GJ, Davis JJ, Disz T, Edwards

RA, Gerdes S, Parrello B, Shukla M, Vonstein V, Wattam AR, Xia F,

Stevens R

. 2014. The SEED and the rapid annotation of microbial ge-

nomes using subsystems technology (RAST). Nucleic Acids Res

42:

D206 –D214.

http://dx.doi.org/10.1093/nar/gkt1226

28.

Aziz

RK, Bartels D, Best AA, DeJongh M, Disz T, Edwards RA,

Formsma K, Gerdes S, Glass EM, Kubal M, Meyer F, Olsen GJ, Olson

R, Osterman AL, Overbeek RA, McNeil LK, Paarmann D, Paczian T,

Parrello B, Pusch GD, Reich C, Stevens R, Vassieva O, Vonstein V,

Wilke A, Zagnitko O

. 2008. The RAST server: rapid annotations using

subsystems technology. BMC Genomics

75.

http://dx.doi.org/10.1186/

1471-2164-9-75

29.

Van

Domselaar GH, Stothard P, Shrivastava S, Cruz JA, Guo A, Dong

X, Lu P, Szafron D, Greiner R, Wishart DS

. 2005. BASys: a web server for

automated bacterial genome annotation. Nucleic Acids Res

33:

W455–W459.

http://dx.doi.org/10.1093/nar/gki593

30.

Markowitz

VM, Chen IM, Palaniappan K, Chu K, Szeto E, Pillay M,

Ratner A, Huang J, Woyke T, Huntemann M, Anderson I, Billis K,

Varghese N, Mavromatis K, Pati A, Ivanova NN, Kyrpides NC

. 2014.

IMG 4 version of the integrated microbial genomes comparative analysis

system. Nucleic Acids Res

42:

D560 –D567.

http://dx.doi.org/10.1093/nar/

gkt963

31.

Markowitz

VM, Chen IM, Chu K, Szeto E, Palaniappan K, Pillay M,

Ratner A, Huang J, Pagani I, Tringe S, Huntemann M, Billis K,

Varghese N, Tennessen K, Mavromatis K, Pati A, Ivanova NN, Kyrpi-

des NC

. 2014. IMG/M 4 version of the integrated metagenome compar-

ative analysis system. Nucleic Acids Res

42:

D568 –D573.

http://

dx.doi.org/10.1093/nar/gkt919

32.

Brooks

SL, Van Hamme JD

. 2012. Whole-genome shotgun sequence of

Rhodococcus

species strain JVH1. J Bacteriol

194:

5492–5493.

http://

dx.doi.org/10.1128/JB.01066-12

33.

Van

Hamme JD, Fedorak PM, Foght JM, Gray MR, Dettman HD

. 2004.

Use of a novel fluorinated organosulfur compound to isolate bacteria

capable of carbon-sulfur bond cleavage. Appl Environ Microbiol

70:

1487–1493.

http://dx.doi.org/10.1128/AEM.70.3.1487-1493.2004

34.

Van

Kessel JC, Hatfull GF

. 2007. Recombineering in

Mycobacterium

tuberculosis

. Nat Methods

147–152.

http://dx.doi.org/10.1038/

nmeth996

35.

YS, Adroub SA, Aleisa F, Mahmood H, Othoum G, Rashid F, Zaher

M, Ali S, Bitter W, Pain A, Abdallah AM

. 2012. Complete genome

sequence of

Mycobacterium fortuitum

subsp.

fortuitum

type strain

DSM46621. J Bacteriol

194:

6337– 6338.

http://dx.doi.org/10.1128/

JB.01461-12

36.

Jones

RJ, Jenkins DE

. 1965. Mycobacteria isolated from soil. Can J Mi-

crobiol

11:

127–133.

http://dx.doi.org/10.1139/m65-018

37.

Covert

TC, Rodgers MR, Reyes AL, Stelma GN, Jr.

1999. Occurrence of

nontuberculous mycobacteria in environmental samples. Appl Environ

Microbiol

65:

2492–2496.

38.

An D, Danhorn T, Fuqua C, Parsek MR

. 2006. Quorum sensing and

motility mediate interactions between

Pseudomonas aeruginosa

and

Agro-

bacterium tumefaciens

in biofilm cocultures. Proc Natl Acad SciUSA

103:

3828 –3833.

http://dx.doi.org/10.1073/pnas.0511323103

39.

Hernandez

ME, Kappler A, Newman DK

. 2004. Phenazines and other

redox-active antibiotics promote microbial mineral reduction. Appl En-

viron Microbiol

70:

921–928.

http://dx.doi.org/10.1128/AEM.70.2.921

-928.2004

40.

Bellin

DL, Sakhtah H, Rosenstein JK, Levine PM, Thimot J, Emmett K,

Dietrich LE, Shepard KL

. 2014. Integrated circuit-based electrochemical

sensor for spatially resolved detection of redox-active metabolites in bio-

films. Nat Commun

3256.

41.

Price-Whelan A, Dietrich LEP, Newman DK

. 2006. Rethinking “second-

ary” metabolism: physiological roles for phenazine antibiotics. Nat Chem

Biol

71–78.

http://dx.doi.org/10.1038/nchembio764

42.

Ramos

I, Dietrich LEP, Price-Whelan A, Newman DK

. 2010. Phenazines

affect biofilm formation by

Pseudomonas aeruginosa

in similar ways at

various scales. Res Microbiol

161:

187–191.

http://dx.doi.org/10.1016/

j.resmic.2010.01.003

43.

Vleesschauwer D, Cornelis P, Höfte M

. 2006. Redox-active pyocya-

nin secreted by

Pseudomonas aeruginosa

7NSK2 triggers systemic resis-

Degradation of Phenazines by Mycobacteria

November/December 2015 Volume 6 Issue 6 e01520-15

®

mbio.asm.org

on January 13, 2016 - Published by

mbio.asm.org

Downloaded from

tance to

Magnaporthe grisea

but enhances

rhizoctonia solani

susceptibility

in rice. Mol Plant Microbe Interact

19:

1406 –1419.

http://dx.doi.org/

10.1094/MPMI-19-1406

44.

Hunter

RC, Klepac-Ceraj V, Lorenzi MM, Grotzinger H, Martin TR,

Newman DK

. 2012. Phenazine content in the cystic fibrosis respiratory

tract negatively correlates with lung function and microbial complexity.

Am J Respir Cell Mol Biol

47:

738 –745.

http://dx.doi.org/10.1165/

rcmb.2012-0088OC

45.

Widdel

F, Kohring G, Mayer F

. 1983. Studies on dissimilatory sulfate-

reducing bacteria that decompose fatty acids. III. Characterization of the

filamentous gliding

Desulfonema limicola

gen. nov. sp. nov., and

Desul-

fonema magnum

sp. nov. Arch Microbiol

134:

286 –294.

http://dx.doi.org/

10.1007/BF00407804

46.

Eder

W, Jahnke LL, Schmidt M, Huber R

. 2001. Microbial diversity of

the brine-seawater interface of the Kebrit Deep, Red Sea, studied via

16S rRNA gene sequences and cultivation methods. Appl Environ Mi-

crobiol

67:

3077–3085.

http://dx.doi.org/10.1128/AEM.67.7.3077

-3085.2001

47.

Eder

W, Ludwig W, Huber R

. 1999. Novel 16S rRNA gene sequences

retrieved from highly saline brine sediments of Kebrit Deep, Red Sea. Arch

Microbiol

172:

213–218.

http://dx.doi.org/10.1007/s002030050762

48.

Letunic

I, Bork P

. 2011. Interactive Tree Of Life v2: online annotation and

display of phylogenetic trees made easy. Nucleic Acids Res

39:

W475–W478.

http://dx.doi.org/10.1093/nar/gkr201

49.

Dereeper

A, Audic S, Claverie J, Blanc G

. 2010. BLAST-EXPLORER

helps you building datasets for phylogenetic analysis. BMC Evol Biol

10:

http://dx.doi.org/10.1186/1471-2148-10-8

50.

Dereeper

A, Guignon V, Blanc G, Audic S, Buffet S, Chevenet F,

Dufayard JF, Guindon S, Lefort V, Lescot M, Claverie JM, Gascuel O

2008. Phylogeny.fr: robust phylogenetic analysis for the non-specialist.

Nucleic Acids Res

36:

W465–W469.

51.

Letunic I, Bork P

. 2007. Interactive Tree Of Life (iTOL): an online tool for

phylogenetic tree display and annotation. Bioinformatics

23:

127–128.

http://dx.doi.org/10.1093/bioinformatics/btl529

52.

Belisle

JT, Mahaffey SB, Hill PJ

. 2009. Isolation of

Mycobacterium

species

genomic DNA, p 1–12. In Parish T, Brown AC (ed), Mycobacteria proto-

cols. Humana Press, New York, NY.

53.

Chin C, Alexander DH, Marks P, Klammer AA, Drake J, Heiner C,

Clum A, Copeland A, Huddleston J, Eichler EE, Turner SW, Korlach J

2013. Nonhybrid, finished microbial genome assemblies from long-read

SMRT sequencing data. Nat Methods

10:

563–569.

http://dx.doi.org/

10.1038/nmeth.2474

54.

Rustad

TR, Roberts DM, Liao RP, Sherman DR

. 2009. Isolation of

mycobacterial RNA, p 13–22.

Parish T, Brown AC (ed), Mycobacteria

protocols, 2nd ed. Humana Press, New York, NY.

55.

Bolger AM, Lohse M, Usadel B

. 2014. Trimmomatic: a flexible trimmer

for Illumina sequence data. BioInformatics

30:

2114 –2120.

http://

dx.doi.org/10.1093/bioinformatics/btu170

56.

Langmead

B, Trapnell C, Pop M, Salzberg SL

. 2009. Ultrafast and

memory-efficient alignment of short DNA sequences to the human ge-

nome. Genome Biol

10:

R25.

http://dx.doi.org/10.1186/gb-2009-10-3-r25

57.

H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, Marth G,

Abecasis G, Durbin R; Genome Project Data Processing

. 2009. The

Sequence Alignment/Map format and SAMtools. Bioinformatics

25:

2078 –2079.

http://dx.doi.org/10.1093/bioinformatics/btp352

58.

Delhomme

N, Padioleau I, Furlong EE, Steinmetz LM

. 2012.

easyRNASeq: a bioconductor package for processing RNA-Seq data.

Bioinformatics

28:

2532–2533.

http://dx.doi.org/10.1093/bioinformatics/

bts477

59.

Goude

R, Parish T

. 2009. Electroporation of mycobacteria, p 203–216.

Parish T, Brown AC (ed), Mycobacteria protocols. Humana Press, New

York, NY.

60.

DasGupta SK, Jain S, Kaushal D, Tyagi AK

. 1998. Expression systems for

study of mycobacterial gene regulation and development of recombinant

BCG vaccines. Biochem Biophys Res Commun

246:

797– 804.

http://

dx.doi.org/10.1006/bbrc.1998.8724

61.

Shanks

RMQ, Caiazza NC, Hinsa SM, Toutain CM, O’Toole GA

. 2006.

Saccharomyces cerevisiae

-based molecular tool kit for manipulation of

genes from gram-negative bacteria. Appl Environ Microbiol

72:

5027–5036.

http://dx.doi.org/10.1128/AEM.00682-06

Costa et al.

®

mbio.asm.org

November/December 2015 Volume 6 Issue 6 e01520-15

mbio.asm.org

on January 13, 2016 - Published by

mbio.asm.org

Downloaded from